EPA-905-R-91-001
Midwest Pollution Control Biologists Meeting
U.S. EPA Region 5
1991
Testing the Toxicity of Field Collected Sediments
Marcia K. Nelson, James J. Coyle and G. Allen Burton
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Nelson, Coyle & Burton MPCB 1991: Sediment Toricity Testing
Course Agenda
TESTING THE TOXICITY OF FIELD COLLECTED FRESHWATER SEDIMENTS"
M.K. Nelson1, JJ. Coyle1 and GA. Burton2
Guest Speaker. R.Wood3
1 U.S. Fish and Wildlife Service
National Fisheries Contaminant Research Center
Columbia, MO
314-875-5399
2 Wright State University
Biological Sciences Department
Dayton, OH
513-873-2201
3 U.S. Environmental Protection Agency
Office of Water Enforcement and Permits
Washington, D.C.
202-475-9534
I. Introduction. Nelson
A. Extent of sediment contamination.
II. Sediment Assessments. Nelson
U.S. Environmental Protection Agency
Region 5, Library (PL- 3 2j)
77 West Jackson Boulevc.j, I2it) Floor
:.:\ '!_ 60604-3590
A. U.S. EPA 1989 Sediment Methods Compendium.
B. ASTM Sediment Sub-committee activities, E47.03.
C. Assessment and Remediation of Contaminated Sediments (ARCS), Great Lakes National
Program Office.
EH. EPA Sediment Management Strategy. Wood
A. Sediment management strategy.
B. Extent.
C. Research driving regulatory solutions.
D. NPDES Program adapting to prevention of sediment contamination.
E. Needs to address sediment contamination prevention.
IV. Safety Precautions and Considerations. Coyle
A. Minimizing exposure. ,,-.._,,.,_
B. Proactive safety management - . ';" J i.'n'
C. Primary, secondary, and tertiary protection
D. Physiological and psychological factors.
E. Perception of hazard. , . ^
F. Route of exposure. • ••'.-• ••.• «J
co
cvj
CD
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V. Sediment Manipulation. Coyle
A. Collection.
B. Shipping.
C. Storage.
D. Preparation.
1. Mixing.
2. Aqueous extractions (i.e., pore water, elutriates).
a. Methods.
b. Practical considerations.
c, Factors influencing composition and toxicity of pore water and elutriates.
E. Water quality.
a. Routine measurements.
b. Potential problems and solutions.
F. Sediment disposal considerations and requirements.
VI. Sediment and aqueous extract chemistry. Nelson
A. Metals and other inorganics.
B. Organics.
VII. Whole sediment characterization. Nelson
A. Total organic carbon.
B. Particle size distribution (percent sand, silt, clay).
C. pH.
D. Total volatile sulfides.
E. Water content (percent).
Sediment Toxicity Testing.
A. Microtox testing of aqueous sediment extractions. Coyle
1. Methods review.
a. Future approaches (Direct Contact).
B. Aqueous extract testing. Burton and Coyle
1. Test organisms (Daphnia mflgpa,, Ceriodaphnia dubia. Pimphales promelas).
2. Methods review.
3. Test set-up.
4. Monitoring test.
5. Ending test.
6. Water Quality.
7. Interpreting results.
a. Tests reflect acute toxicity of water soluble contaminants.
b. Tests results not stand-alone descriptions, but are parts of a larger toxicity appraisal
process.
C. Microbial and In situ Testing. Burton
D. Whole Sediment Testing. Nelson and Burton
L Initiating tCStS.
a. Experimental design.
2. Test organisms (Hyalella azteca. Chironomus riparius. C^jr^nomus tentans. Daphnia
Ceriodaphnifl dubia).
a. Culture.
b. Handling.
c. Test preparations.
(1) Diluter calibration.
(2) Food preparation.
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Nelson, Coyle & Burton MPCB 1991: Sediment Toxitity Testing Course Agenda
(3) Temperature in water bath.
3. Test set-up.
a. Day -1.
(1) Sediment into test chambers.
(2) Overlying water.
(3) Aeration.
b. Day 0.
(1) Water quality determinations.
4. Monitoring tests.
a. Biological
(1) Feeding.
(2) Qualitative observations.
(a) Test organisms.
(b) Sediment and overlying water conditions.
b. Equipment operation.
(1) Diluter functioning.
(2) Aeration.
(3) Screens cleaned.
c. Water quality determinations.
(1) Day 7, etc. to end of test.
5. Ending tests.
a. Water quality.
b. Sieving sediments.
c. Retrieving test organisms.
d. Preserving test organisms.
6. Interpreting results.
a. Test acceptability.
DC Strengths and Limitations of Sediment Toxicity Testing. OPEN FORUM
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United States June 1989
Environmental Protection
Agency
Watersneo Protection Division
Final
Report
Classification
Methods Compendium
Mike Kravitz
U.S. EPA
Office of Water Regulations and Standards
(WH-553)
401 M. St. S.W.
Washington, B.C. 20460
202-475-8085
FTS 475-8085
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Draft Final Report
SEDIMENT CLASSIFICATION
METHODS COMPENDIUM
by
U.S. Environmental Protection Agency
Portions of this document were prepared by
Tetra Tech, Inc., under the direction of
Michael Kravitz, U.S. EPA Work Assignment Manager
June 1989
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CONTENTS
Page
LIST OF FIGURES ix
.1ST CF TABLES x
-CKNOWLEDGMENTS xi
CHAPTER 1. INTRODUCTION 1-1
1.0 BACKGROUND 1-1
2.0 OBJECTIVE 1-2
3.0 OVERVIEW 1-2
CHAPTER 2. BULK SEDIMENT TOXICITY TEST APPROACH 2-1
1.0 SPECIFIC APPLICATIONS 2-1
1.1 Current Use 2-1
1.2 Potential Use 2-2
2.0 DESCRIPTION 2-3
2.1 Description of Method 2-3
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection 2-7
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 2-7
3.0 USEFULNESS 2-8
3.1 Environmental Applicability 2-8
3.2 General Advantages and Limitations 2-10
4.0 STATUS 2-13
4.1 Extent of Use 2-13
4.2 Extent to Which Approach Has Been Field-Validated 2-13
4.3 Reasons for Limited Use 2-13
4.4 Outlook for Future Use and Amount of Development Yet
Needed 2-13
5.0 REFERENCES 2-14
11
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CHAPTER 3. SPIKED-SEDIMENT TQXICITY TEST APPROACH 2-1
1.0 SPECIFIC APPLICATIONS 3-1
1.1 Current Use 3-1
1.2 Potential Use 3-2
2.0 DESCRIPTION 3-2
2.1 Description of Method 3-2
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection 3-6
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 3-7
3.0 USEFULNESS 3-8
3.1 Environmental Applicability 3-8
3.2 General Advantages and Limitations 3-10
•l.O STATUS 3-13
4.1 Extent of Use 3-13
4.2 Extent to Which Approach Has Been Field-Validated 3-13
4.3 Reasons for Limited Use 3-14
4.4 Outlook for Future Use and Amount of Development Yet
Needed 3-14
5.0 REFERENCES 3-14
CHAPTER 4. INTERSTITIAL WATER TOXICITY APPROACH 4-1
1.0 SPECIFIC APPLICATIONS 4-1
1.1 Current Use 4-1
1.2 Potential Use 4-2
2.0 DESCRIPTION 4-2
2.1 Description of Method 4-2
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection 4-16
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 4-16
3.0 USEFULNESS 4-17
3.1 Environmental Applicability 4-17
3.2 General Advantages and Limitations 4-19
111
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4.0 STATUS 4-21
4.1 Extent of Use _ 4-21
4.2 Extent to Which Approach Has Been Field-Validated 4-22
4.3 Reasons for Limited Use 4-22
4.4 Outlook for Future Use and Amount of Development Yet
Needed 4.22
5.0 REFERENCES 4.23
CHAPTER 5. EQUILIBRIUM PARTITIONING APPROACH 5-1
1.0 SPECIFIC APPLICATIONS 5-1
1.1 Current Use 5-2
1.2 Potential Use 5.3
2.0 DESCRIPTION 5-4
2.1 Description of Method 5-4
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection ' 5.7
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 5.3
3.0 USEFULNESS 5-9
3.1 Environmental Applicability 5.9
3.2 General Advantages and Limitations 5-11
4.0 STATUS 5-15
4.1 Extent of Use 5-16
4.2 Extent to Which Approach Has Been Field-Validated 5-16
4.3 Reasons for Limited Use 5-17
4.4 Outlook for Future Use and Amount of Development Yet
Needed 5.17
5.0 DOCUMENTS 5-18
CHAPTER 6. TISSUE RESIDUE APPROACH 6-1
1.0 SPECIFIC APPLICATIONS 6-2
1.1 Current Use 6-2
1.2 Potential Use 6-2
2.0 DESCRIPTION 5.3
2.1 Description of Method 6-3
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection 6-9
iv
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2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 6-10
3.0 USEFULNESS 6-10
3.1 Environmental Applicability 6-10
3.2 General Advantages and Limitations 6-14
4.0 STATUS 6-17
4.1 Extent of Use 5-17
4.2 Extent to Which Approach Has Been Field-Validated 6-17
4.3 Reasons for Limited Use 6-18
4.4 Outlook for Future Use and Amount of Development Yet
Needed 6-18
5.0 REFERENCES 6-19
CHAPTER 7. FRESHWATER BENTHIC MACROINVERTEBRATE COMMUNITY STRUCTURE
AND FUNCTION 7-1
1.0 SPECIFIC APPLICATIONS 7-2
1.1 Current Use 7-2
1.2 Potential Use 7-5
2.0 DESCRIPTION 7-6
2.1 Description of Method 7-6
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection 7-28
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 7-28
3.0 USEFULNESS 7-28
3.1 Environmental Applicability 7-28
3.2 General Advantages and Limitations 7-30
4.0 STATUS 7-35
4.1 Extent of Use 7-35
4.2 Extent to Which Approach Has Been Field-Validated 7-35
4.3 Reasons for Limited Use 7-36
4.4 Outlook for Future Use and Amount of Development Yet
Needed 7-36
5.0 REFERENCES 7-36
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CHAPTER 8. MARINE BENTHIC COMMUNITY STRUCTURE ASSESSMENT 8-1
1.0 SPECIFIC APPLICATIONS 8-2
1.1 Current Use 3.3
1.2 Potential Use 8-7
2.0 DESCRIPTION 3.3
2.1 Description of Method 3.3
2.2 Applicability of Method to Human Health, Aquatic Life,
or wildlife Protection 8-20
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 8-21
3.0 USEFULNESS 8-21
3.1 Environmental Applicability 3-22
3.2 General Advantages and Limitations 8-26
4.0 STATUS 3-31
4.1 Extent of Use 8-31
4.2 Extent to Which Approach Has Been Field-Validated 8-32
4.3 Reasons for Limited Use 8-32
4.4 Outlook for Future Use and Amount of Development Yet
Needed 8-32
5.0 REFERENCES 3.34
CHAPTER 9. SEDIMENT QUALITY TRIAD APPROACH 9-1
1.0 SPECIFIC APPLICATIONS 9-1
1.1 Current Use 9-1
1.2 Potential Use 9-2
2.0 DESCRIPTION 9-2
2.1 Description of Method 9-2
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection 9-15
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 9-16
3.0 USEFULNESS 9-16
3.1 Environmental Applicability 9-16
3.2 General Advantages and Limitations 9-20
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4.0 STATUS 9-24
4.1 Extent of Use 9-24
4.2 Extent to Which Approach Has Been Field-Validated 9-24
4.3 Reasons for Limited Use 9-24
4.4 Outlook for Future Use and Amount of Development Yet
Needed 5
5.0 REFERENCES^ 9-25
CHAPTER 10. APPARENT EFFECTS THRESHOLD APPROACH 10-1
1.0 'ECIFIC APPLICATIONS •"-!
1.1 Current Use 10-1
1.2 Potential Use 10-4
2.0 DESCRIPTION 10-5
2.1 Description of Method 10-5
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection 10-16
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 10-16
3.0 USEFULNESS 10-17
3.1 Environmental Applicability 10-17
3.2 General Advantages and Limitations 10-22
4.0 STATUS 10-33
4.1 Extent of Use 10-33
4.2 Extent to Which Approach Has Been Field-Validated 10-35
4.3 Reasons for Limited Use 10-37
4.4 Outlook for Future Use and Amount of Development Yet
Needed 10-37
5.0 REFERENCES 10-38
CHAPTER 11. A SUMMARY OF THE SEDIMENT ASSESSMENT STRATEGY RECOMMENDED
BY THE INTERNATIONAL JOINT COMMISSION 11-1
1.0 SPECIFIC APPLICATIONS 11-1
1.1 Current Use 11-1
1.2 Potential Use 11-2
vn
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2.0 DESCRIPTION 11-2
2.1 Description of Method 11-2
2.2 Applicability of Method to Human Health, Aquatic Life,
or Wildlife Protection 11-14
2.3 Ability of Method to Generate Numerical Criteria for
Specific Chemicals 11-14
3.0 USEFULNESS 11-15
3.1 Environmental Applicability 11-15
3.2 General Advantages and Limitations 11-16
4.0 STATUS 11-19
4.1 Extent of Use 11-19
4.2 Extent to Which Approach Has Been Field-Validated 11-19
4.3 Reasons for Limited Use 11-20
4.4 Outlook for Future Use and Amount of Development Yet
Needed 11-20
5.0 REFERENCES 11-20
VI 1 1
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FIGURES
Number Page
4-1 Overview of the Phase I toxicity characterization process 4-7
9-1 Conceptual model of the Sediment Quality Triad 9-3
9-2 Triaxial plots of eight possible outcomes for Sediment
Quality Triad results 9-14
10-1 The AET approach applied to sediments tested for lead and
i-methylphenol concentrations and toxicity response during
Dioassays 10-7
10-2- Measures of reliability (sensitivity and efficiency) 10-31
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TABLES
1-1 Sediment quality assessment methods 1-3
1-2 Structure of sediment quality assessment method chapters 1-6
4-1 Phase I characterization results and suspect toxicant
classification for two effluents 4-12
9-1 Current uses of the Sediment Quality Triad approach 9-4
9-2 Possible conclusions provided by using the Sediment Quality
Triad approach 9-6
9-3 Example analytes and detection limits for use in the
chemistry component of Triad 9-9
9-4 Possible static sediment bioassays 9-11
10-1 Selected chemicals for which AET have been developed in
Puget Sound 10-18
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ACKNOWLEDGMENTS
This compendium was prepared by the U.S. Environmental Protection Agency,
Sediment Oversight Technical Committee. Chaired by Or. Elizabeth Southerland
of the Office of Water Regulations and Standards, the committee has represen-
tation from a number of Program Offices in Headquarters and the Regions.
The methods represented here were written by the following authors (also
listed at the beginning of their respective chapters):
• Gerald Ankley, Anthony R. Carlson, Phillip M. Cook, Wayne S.
Davis, Catherine Krueger, Janet Lamberson, Henry Lee II,
Richard C. Swartz, Nelson Thomas, and Christopher S. Zarba
(U.S. EPA)
• Gordon R. Bilyard, Gary M. Braun, and Betsy Day (Tetra Tech,
Inc.)
• Peter M. Chapman (E.V.S. Consultants, Ltd.)
• Philippe Ross (Illinois Natural History Survey)
• Joyce E. Lathrop (Stream Assessments Company).
Critical reviews of portions of this document were provided by the following
U.S. EPA persons: Gerald Ankley, Carol Bass, Dave Cowgill, Philip Crocker,
Shannon Cunniff, Kim Devonald, Cynthia Fuller, Ray Hall, David Hansen,
Nicholas Loux, Menchu Martinez, Brian Melzian, Ossie Meyn, James Neiheisel,
Dave Bedford, Greg Schweer, Richard Swartz, Nelson Thomas, Mark Tuchman,
Gerald walsh, Al Wastler, Howard Zar, and Chris Zarba.
Assistance in preparation and production of the compendium was provided
by retra Tech, Inc. in partial fulfillment of EPA Contract No. 68-03-3475.
Dr.
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TABLE 1-1. SEDIMENT QUALITY ASSESSMENT METHODS
(Sediment Classification Methods Compendium, U.S. EPA, June 1989)
Method (Chapter)
Nura Descr Comb
Concept
Bulk Sediment Toxicity
(2.0)
Test organisms are exposed to sediments which may contain unknown quantities of
potentially toxic chemicals. At the end of a specified time period, the response of
the test organisms is examined in relation to a specified biological endpomt.
Spiked Sediment Toxicity
(3.0)
Dose-response relationships are established by exposing test organisms to
sediments that have been spiked with known amounts of chemicals or mixtures of
chemicals.
Interstitial Water Toxicity
(4.0)
Toxkityof interstitial water isquantified and identification evaluation procedures
are applied to identify and quantify chemical components responsible for sediment
toxicity. The procedures are implemented in three phases to characterize
interstitial water toxicity, identify the suspected toxicant, and confirm toxicant
identification.
Equilibrium Partitioning
(5.0)
A sediment quality value for a given contaminant is determined by calculating the
sediment concentration of the contaminant that would correspond to an interstitial
water concentration equivalent to the U.S. EPA water quality criterion for the
contaminant.
Tissue Residue
(6.0)
Safe sediment concentrations of specific chemicals are established by determining
the sediment chemical concentration that will result in acceptable tissue residues
Methods to derive unacceptable tissue residues are based on chronic waterqualiry
criteria and bkxoncentration factors, chronic dose-response experiments or field
correlation, and human health risk levels from the consumption of freshwater fish
or seafood.
Freshwater Benthic Community Structure
(8.0)
Environmental degradation is measured by evaluating alterations in freshwater
benthic community structure.
Marine Benthic Community Structure
(9.0)
Environmental degradation is measured by evaluating alterations in manne benthic
community structure.
Sediment Quality Triad
(9.0)
Sediment chemical contamination, sediment toxicity, and benthic infauna
community structure are measured on the same sediment. Correspondence
between sediment chemistry, toxicity, and biological effects is used to determine
sediment concentrations that discriminate conditions of minimal, uncertain, and
major biological effects.
Apparent Effects Threshold
(10.0)
An AET is the sediment concentration of a contaminant above which statistically
significant biological effects (e.g-, amphipodmortality in bioassays, depressions in
the abundance of benthic infauna) would always be expected. AET values are
empirically derived from paired field data for sediment chemistry and a range of
biological effects indicators.
International Joint Commission
(11.0)1
Contaminated sediments are nttrttrrt in two stages: 1) an initial assessment that
is based on macro-zoobenthk community structure and concentrations of
contaminants in sediments and biological tissues, and 2) a detailed assessment that
» based on a phased sampling of the physical, chemical, and biological aspects of
the sediment, including laboratory toxicity bioassays.
The LIC approach is an example of a sequential approach, or 'strategy* combining a number of methods for the purpose of
contaminated sediments in the Great Lakes.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM Sediment Subcommittee
American Society for Testing and Materials
E-47 Biological Effects and Environmental Fate (Main Committee)
E-47.03 Sediment Toxicity Subcommittee
Christopher G. Ingersoll, Chair
USFWS, NFCRC
Columbia, MO, 314/875-5399
ASTM Sediment Subcommittee Activities
Document #1: E 1383 Guide for Conducting Sediment Toxicity Tests with Freshwater Invertebrates (Task
Group Chair: Marcia Nelson, NFCRC, Columbia, MO, 314/875-5399).
Proposed additional species-specific annexes.
(1) Daphnia and Ceriodaphnia (Allen Burton, Wright State University, Dayton, OH, 513/873-2201).
(2) Diporeia spp. (formerly Pontoporeia hovi: Peter Landrum, NOAA, Ann Arbor, MI, 313/668-2276).
(3) Ostracods (Arthur Stewart, Oak Ridge National Laboratory, Oak Ridge, TN, 615-574-7835).
(4) Hexaeenia spp. (Donna Bedard, Ontario Ministry of the Environment, Rexdale, Ontario, 416/235-5970
and Mary Henry, USFWS, U. of Minn, Minneapolis, MM).
(5) Tubificid oligochaetes (Trefor Reynoldson, Environment Canada, Burlington, Ontario, 416/336-4783).
(6) Naidid oligochaetes (Dave Smith, Bio-Aquatics Testing, Carrollton, TX, 214/247-5928).
(7) Lumbricus sp. (Gary Phipps, ERL-Duluth, MN, 218/720-5550).
(8) Mollusks (Don Wade and Anne Keller, TVA, Muscle Shoals, AL, 205/386-2068).
Document #2: E 1367 Guide for Conducting 10-d Static Sediment Toxicity Tests with Estuarine and Marine
Amphipods (Task Group Chair: Janet Lamberson, USEPA, Newport, OR, 503/867-4043).
Document #3: E 1391 Guide for Collection, Storage, Characterization, and Manipulation of Sediment for
Toxicological Testing (Task Group Chair: A. Burton, WSU).
Document #4: Guide For Designing. Sediment Toxicity and Bioaccumulation Tests (Task Group Chair: John
Scott, SAIC, Narragansett, RI, 401/782-3017).
Document #5: Sediment Resuspension Testing Methods (Allen Burton, WSU).
Document #6: Guide for Conducting Sediment Toxicity Tests with Polychaetes (Task Group Chair: Don
Reish, California State University-Long Beach, Long Beach, CA, 213/431-7064).
Document #7: Guide for Determination of the Bioaccumulation of Sediment-Associated Contaminants by
Fish (Draft #2, 04/17/90, Task Group Chair: Mike Mac, USFWS, Ann Arbor, MI, 313/994-3331).
Document #8: Guide for Determination of the Bioaccumulation of Sediment-Associated Contaminants by
Benthic Invertebrates. (Task Group Chair: Henry Lee, USEPA, Newport, OR, 503/867-4042).
Document #9: Use of Oysters and Echinoderm Embryos and Larvae in Sediment Toxicity Testing (Task
Group Chair: Paul Dinnel, University of Washington, Seattle, WA, 206/543-7345).
Document #10: Toxicity Identification and Evaluation (TIE) for Sediment Water Extracts (Task Group
Chair: vacant).
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Robert Wood, USEPA
SEDIMENT MANAGEMENT: A REGULATORY PERSPECTIVE
Robert Wood
U.S. Environmental Protection Agency
Office of Water Enforcement and Permits
401 M St., S.W.
Washington, D.C. 20460
202-475-8488
I. Introduction.
A. Topics to be covered.
1. EPA agency-wide sediment management strategy.
2. Why a strategy now?
a. Extent of sediment contamination problem.
b. What we have learned through research that is driving regulatory solutions.
3. How EPA envisions NPDES Program adapting to address prevention of sediment
contamination caused by point sources.
4. What do we need (research, procedures, policy) in order for the NPDES Program
to address sediment contamination prevention?
II. EPA sediment management strategy.
A. The strategy will state EPA's policy on sediments in light of latest science and understanding
of the extent of the problem. It is very early in the strategy development process. EPA is
committed to involving the public in the process.
B. The strategy will likely have 4 basic components.
1. Assessment and risk identification.
a. Statement of the sediment contamination problem, why we think its a
national problem, how we know it is a problem in some locations.
b. What EPA intends to do to better define the extent of the national
problem.
2. Prevention.
a. Statement of policy on point and non-point source prevention, pesticide
regulation, and toxic substances control.
3. Remediation.
a. Roles and responsibilities.
b. Consistent identification of sites for remediation.
c. Consistent cleanup goals.
4. Dredged material management.
a. Balancing economic and environmental factors.
b. Applicability of RCRA.
m. Why a sediment management strategy now?
A. What data is telling us about risk and ecological impact.
1. 1989 National Academy of Sciences Report on contaminated marine sediments.
2. Site-specific studies showing human health risk from consumption of fish and
shellfish.
a. Quincy Bay, MA: cancer risk from consuming lobster tomalley.
b. Lake Michigan: developmental problems in children whose mothers
consumed large amounts of fish.
c. Los Angeles-Long Beach Harbor 10"3-"4 cancer risk from consuming white
croaker.
d. Puget Sound: As much as 2 x 10"4 cancer risk for moderate seafood
consumers and 4 x 10~3 risk for high-quantity consumers.
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3. Site-specific studies showing harm to aquatic life, waterfowl, and up the food chain.
a. Elizabeth River, VA: Severe fin and gill erosion, tumors, and mortality.
b. Black River, OH: fish tumors.
c. Great Lakes: reproductive problems in Forster's tern, reproductive failures
and mortality in mink.
d. Commencement Bay, WA: mortality in amphipods and oyster larvae.
B. Improved ability to identify sediment toxicity and classify sediments based on their impact on
aquatic life and human health.
1. Criteria documents.
a. Scheduled for public review and comment in 1991. (6 non-polar organics).
2. Advances in whole sediment toxicity tests.
3. Advances in sediment TIE research and method development making TIE
methodologies increasingly useful for identifying causative agents and sources.
C. Congress is interested. Seven separate pieces of legislation introduced in 89 and 90 that
address sediments.
1. National inventory of sites.
2. Sediment criteria and standards.
3. Accelerated point and non-point source controls.
IV. NPDES Program
A. EPA fully intends to use sediment criteria, sediment toxicity analysis, and sediment TIE as
the basis for point source controls to protect sediment quality.
1. EPA believes the science of sediment classification and source identification is solid
and getting better and that implementing point source controls will therefore not
require any great leap of faith.
B. What is on the horizon. Point source sediment quality controls are probably inevitable.
1. Source identification using refined sediment TIE procedures.
2. Chemical-specific permit limits based on sediment quality criteria.
3. Whole effluent limits based in some way on ambient sediment toxicity (measured or
projected).
4. Chemical-specific permit limits based in the presence of bioconcentratable
compounds on effluent, ambient sediment and/or ambient tissue (measured or
projected).
V. How NPDES gets from here to there.
A. Assessment needs.
1. We know a good deal about the extent of sediment contamination, but we need
more and better information, particularly on source identification.
2. EPA is wrestling with the assessment question. How extensive should an
assessment be?
a. Data base of existing information on sites?
b. Fill in gaps in existing data on sites?
c. Full blown comprehensive assessment (new data) on sites and sources.
B. Need to continue sediment criteria development.
1. First set of 6 non-polar organics.
2. Metals.
3. More organics, inorganics.
C. Need to continue refinement of TIE methodologies.
1. Research so far has been mostly on identifying causative agents in highly complex
sediments. Upcoming research will focus also on less complex samples with
defensible source identification as an objective. EPA is currently selecting
candidate sediment samples for this purpose.
D. Continued refinement of promising sediment toxicity protocols that are user friendly and
suitable for wide use by regulatory authorities.
E. Simplified models of sediment fate and transport that are user friendly and suitable for wide
use by regulatory authorities.
F. Validation
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Robert Wood, USEPA
1. Audience can appreciate the need for validation of predictive methodologies to
show that whatever the methodology, it is reasonably accurate at projecting and
defining real aquatic life and human health risk.
2. EPA is committed to basing point source sediment quality controls in good solid
science. Want to target regulatory efforts at real problems.
G. Need input from scientific community, regulators, and industry. There will be key
opportunities for this.
1. Public comment on agency-wide sediment management strategy (early 1991).
2. Public comment on proposed sediment criteria for 6 non-polar organics (1991).
3. Continued exchanges like today.
VI. Summary.
A. There is strong momentum toward point source sediment contamination controls.
B. In an atypical fashion, the research is driving policy and the regulatory program. This is a
good thing that is likely to yield informed, fair regulatory decisions.
C. We are at a point where we know we are on the right track technically.
D. EPA focus will continue to be on refining methodologies in order to make point source
sediment contamination controls real
YD. Questions.
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For Information on EPA Sedjpieqt Management Strategy Contact:
Betsy Souther-land
U.S. EPA Office of Water Regulations and Standards (WH-553)
401 M St., S.W.
Washington, D.C. 20460
phone: 202-382-7046
FTS 382-7046
T«'T! Wall
U.o. EPA Office of Water Regulations and Standards (WH-553)
401 M St., S.W.
Washington, D.C. 20460
phone: 202-382-7037
FTS 382-7037
For Information on EPA Sediment Criteria Development Contact:
Christopher Zarfoa
U.S. EPA Office of Water Regulations and Standards (WH-585)
401 M St., S.W.
Washington, D.C. 20460
phone: 202-475-7326
FTS 475-7326
For Information on EPA Sediment TIE Research Contact:
Gary Ankley
U.S. EPA Environmental Research Lab
6201 Congdon Blvd.
Duluth,MN 55804
phone: 218-720-5603
FTS 780-5603
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Sediment Safety Procedures
SEDIMENT STORAGE, HANDLING AND TESTING PROCEDURES,
FISH AND INVERTEBRATE TOXICOLOGY
I. General:
This SOP describes the procedures to minimize exposure of personnel and the facility while conducting
laboratory tests with sediments or sediment extracts. Sediment is often a storage reservoir for many
contaminants introduced into surface waters. These contaminants may include polychlorinated biphenyls,
polynuclear aromatic compounds and inorganic contaminants including heavy metals. Contaminants present
in sediment may include carcinogens, mutagens, or potentially toxic compounds. Bioassessment tests (toxicity
and bioaccumulation) are used to estimate potential biological impact that may result from exposure to these
contaminants associated with sediment Since field sediments may contain potentially toxic materials they
should be treated with caution to minimize occupational exposure to workers.
H. Safety:
A. Site Section: Prior to collection of sediment for laboratory tests, information on known or
suspected contaminants associated with the sediment at the site must be identified.
Historical data (e.g., types of industry, known contaminant inputs, STORET) or additional
chemical analyses will be needed before sediments are collected for laboratory tests.
B. Personal protection: This section deals with the procedures that will be implemented by all
personnel working with contaminated sediment. It should be noted that research conducted
with sediment varies considerably depending on the scope and objective of the research.
Therefore, the guidelines set forth in this SOP may not be applicable to all situations dealing
with potentially contaminated sediments (1,23,4).
1. Medical Surveillance. Health monitoring will be provided for personnel working
with sediments. The health monitoring establishes a baseline to which all
subsequent medical finding can be compared.
2. Personal precautions. Workers must always be aware of possible points of
contamination as described by the supervisor. Hands should always be kept away
from the eyes and mouth. After completion of a manipulation involving sediment
or the removal of possibly contaminated laboratory clothing (gloves, lab coat, etc.),
the hands, forearms, and other areas of suspected contact should be washed with
hand soap and water at a sink located within the laboratory work area. Do not use
organic solvents to clean the skin. These solvents may increase penetration of the
contaminant into the skin.
3. Laboratory clothing. When working with sediments it is of the utmost importance
to avoid skin contact A fully fastened knee length lab coat must be worn in the
laboratory work area at all times. Disposable Tyvec" lab clothing must be worn for
sediment manipulation and when water quality is determined. Cloth lab clothing
may be worn during non-hazardous activities, such as feeding test organisms,
entering data, or checking diluters. Any laboratory clothing containing holes or
tears will not be used. The lab coat must be removed and stored in the proper bag
prior to leaving the laboratory work area. All lab clothing may only be handled
while wearing gloves. The procedure for putting on gloves and a lab coat is: (a) put
on one pair of clean gloves, (b) put on the lab coat, and (c) put on a second pair of
gloves. The procedure for removing the gloves and lab coat is: (a) remove the
outer pair of gloves making sure not to contact the skin with the surface of the
outer glove, (b) remove the lab coat, (c) remove the second pair of gloves, and (d)
-------
wash hands at the sink. Clothing should be examined daily for possible
contamination.
4. Hand protection. Hands will be the most frequent point of potential contact with
contaminants. Gloves must be worn to avoid skin contamination. Disposable
gloves must be discarded after each use in appropriate containers designated for
this use. Double gloves will be used with the outer glove being striped off after any
potential exposure. Torn or punctured gloves must be discarded and replaced
immediately. It must be remembered that rubber, latex or vinyl gloves do not
provide full protection. Contaminants may diffuse into the gloves. When sediment
is handled gloves should be changed frequently (3). Cuffs must be tight fitting or
taped to the sleeve to prevent inward migration of contaminants.
5. Eye protection. Safety glasses must be worn at all times. In addition, face shields
will be made available in the laboratory work area.
6. Further precautions. Protective disposable footwear is recommended during
sediment manipulation. Long hair should be tied back and loose clothing should be
covered by the lab coat. Eating, drinking, smoking, chewing gum, smokeless
tobacco and shorts are prohibited in the laboratory work area where sediments are
being used or stored. Food must not be stored in the laboratory work area. Oral
pipetting will never be performed. In addition, respirators, a glove box, or a vented
hood will be used when sediment is manipulated. Respirators will be labeled with
the workers name, date of filter replacement and stored in individual lockers when
not in use. These lockers are located in the change area outside the laboratory
work area. Reusable protective gear will be placed in a cabinet located outside the
laboratory work area (see Section C below).
C. Facility engineered protection: The following guidelines are for the laboratory work area
where sediments will be tested.
1. Area identification and access control.
a. The laboratory work area where sediments are used or stored will be
properly identified. A sign stating "Authorized personnel only" will be
visible. Access to the designated laboratory work area will be limited.
Access doors to the building will be kept dosed while sediment is
manipulated.
c. Animals and plants not related to the experiment shall not be permitted in
the laboratory.
2, Eyewash stations and hand washing facilities are available in the laboratory work
area.
3. Containment devices. Work with sediment will be performed in an appropriate
containment device. Procedures involving sediment will not be conducted on an
open bench due to the potential hazard of generating contaminated dusts, aerosols,
or fumes. Hoods, glove boxes, and enclosed vented water baths for testing are used
to minimiM-. the worker exposure to contaminants associated with sediment. All
containment devices will be constructed out of smooth, unbreakable material, such
as TeflonR, stainless steel, polyethylene, fiberglass, or plexiglass. Exhaust air from
hoods, glove boxes, or water baths which contain sediments does not have to be
filtered (1). The discharge must be out of the building, as far from the air intake
supply as possible (1).
4. Equipment. Use of instruments such as pH, dissolved oxygen or conductivity
meters will be used in a glove box or hood. This equipment will be enclosed in
plastic to reduce the potential for contamination. Instruments will be serviced or
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Sediment Safety Procedures
calibrated in the work area. All calibration and maintenance log books should be
kept with the equipment. All equipment that has come in contact with potentially
contaminated sediment must be kept either under negative pressure (e.g., a hood)
or sealed in an air tight container (e.g., a Tupperware" container) before it is
cleaned
5. Work surfaces. All work surfaces potentially exposed to sediments must be covered
with Teflon" sheets, plastic trays, dry absorbent plastic-backed paper, foil, or other
impervious or disposable material. If a surface becomes contaminated or if a spill
occurs, the work surface should be decontaminated or disposed of immediately.
6. Housekeeping. The laboratory work area shall be kept dean and orderly. Clean-
up shall follow every operation or, at a minimum, at the end of each day.
Containers for disposal of contaminated materials will be placed in the work area.
7. Spill control. A sediment spill will be treated as a "Chemical Spill: Organic solvent."
The sediment spill will be contained with the appropriate absorbent material. If a
spill occurs the worker should (a) pour absorbent material on the spill quickly,
using enough material to adsorb all fluid and cover the mass with excess dry
absorbent to control vapors; (b) sound the air horn to signal for help if necessary,
(c) close doors to all labs in the building; (d) increase ventilation by turning on
exhaust hoods in the laboratory work area; (e) if problems are encountered in
containing the spill, consideration should be given to evacuating the building, route
personnel away from the problem area; (f) clean up adsorbents and dispose of them
properly, (g) allow personnel to return to the laboratory work area.
HI. Storage of sediment:
A. Solid-phase sediment and sediment extracts will be stored at 4°C in air-tight containers in
the dark. All samples must be accompanied with proper identification and sample tracking
information. Sediment extracts can be temporarily stored at 4°C in refrigerators located in
the laboratory work areas.
IV. Homogenization and preparation of elutriate samples:
A. Sediment will always be transferred using double containment. Transfer of sediment from
the storage container is a procedure which involves a potential hazard for personal
contamination. During this procedure, the number of investigators in the laboratory work
area should be minimized Other workers in the building must be notified of the handling
of the sediment.
B. Mixing and sampling of solid-phase sediment or sediment extracts will be done in the
original storage container under a hood If the containers holding sediment are removed
from the hood an intermediate non-breakable container must be used. The worker must
use a respirator with organic vapor-acid gas filters and appropriate clothing as described in
Section II when solid-phase sediment or sediment extracts are not under a hood or in a
glove box.
V. Placing sediment (or sediment extracts) into test chambers:
A. Sediment will always be transferred using double containment. Sediment transfer into test
chambers is a procedure which involves a potential hazard for personal contamination.
During this procedure, the number of investigators in the laboratory work area should be
minimized. Other workers in the building must be notified of the handling of the sediment.
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B. Solid-phase sediment will be distributed into the test chambers using a spoon within the
glove box or hood located in the laboratory work area. Mixing and sampling of solid-phase
sediment ing will be done in the original storage container. An aliquot of the solid-phase
sediment is added to each test chamber using a spoon. The solid-phase sediment aliquot in
the test chamber is settled by smoothing with a spoon. Overlying water is place over the
sediment for the test chamber is removed from the hood. Sediment extracts will always be
handled under a hood. When the test chambers are removed from the glove box hood, or
water bath, an intermediate non-breakable container must be used. The worker must use a
respirator with organic vapor-acid gas filters and appropriate clothing as described in Section
II when test chambers containing solid-phase sediment or sediment extracts are not under
the vented water bath, hood, or in a glove box.
VI. Conducting
A. Hoods or incubators will be used to manipulate and solid-phase sediment and sediment
extracts.
B. Water baths are covered with a vented plexiglass hood. These hoods will only be opened
when: (1) transferring test chambers in and out of the water bath, (2) placing animals into
the test chambers to start a test, (3) feeding the animals, or (4) during water sampling.
VII. Terminating sediment tests:
A. Removal of sediment containing test chambers from plexiglass vented hoods is a procedure
which involves a potential hazard for personal and surface contamination. The number of
investigators in the laboratory work area should be minimized. If the test chambers are
removed from the glove box hood, or water bath, an intermediate non-breakable container
must be used.
B. The worker will use a respirator and appropriate clothing as described in Section II during
transfer of sediment test chambers to the glove box or hood. Sediments may need to be
sieved to enumerate and observe animals.
C. All test chambers and equipment coming in contact with the sediment will be rinsed of
excess sediment in the glove box or hood.
Vin. Clean-up of equipment after sediment tests:
A. Glassware and equipment coming in contact with sediment will be cleaned as soon as
possible. Cleaning glassware poses an increased exposure hazard, all glassware must be
cleaned under the vented sinks or hoods located in the laboratory work area.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Sediment Safety Procedures
References:
1. Dornhoffer, M.K. 1986. Handling chemical carcinogens: A safety guide for the Laboratory
Researcher. Chemical Sciences Laboratories, Lenexa, KS. 62 p.
2. Federal Register, Vol. 43(247):60109-60129.
3. Castegnaro, M. and EJ3. Sansone. 1986. Chemical Carcinogens. Some Guidelines for handling and
disposal in the laboratory. Springer-Verlag, New York. 97 p.
4. Prudent Practices for Disposal of Chemicals from Laboratories. 1983. Committee on Hazardous
Substances in the Laboratory. Commission on Physical Sciences, Mathematics, and Resources.
National Research Council National Academy Press, Washington D.C. 282 p.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1391 (in press)
Draft # 6
June 1990
This document is in process of development and is for ASTM
committee use only. It shall not be reproduced or circulated or
quoted, in whole or in part, outside of ASTM committee activities
except with the approval of the chairman of the committee having
jurisdiction or the President of the Society.
STANDARD GUIDE FOR COLLECTION, STORAGE, CHARACTERIZATION,
AND MANIPULATION OF SEDIMENTS
FOR TOZICOLOGICAL TESTING
G. Allen Burton1 and Peter F. Landrum2
1. Scope
1.1 This guidance document describes procedures for obtaining, storing,
characterizing, and manipulating saltwater and freshwater sediments, for use in
laboratory sediment toxicity evaluations. It is not meant to provide guidance
for all aspects of sediment assessments, such as chemical analyses or monitoring
geophysical characterization, or extractable phase/fractionation analyses. Some
of this information might, however, have applications for some of these
activities, for guidance on toxicity test design and exposure method
considerations, see Guide for Designing Biological Tests with Sediments (Draft
#2) or specific sediment toxicity test methods, (see Section 2.1).
Methodological considerations which affect toxicity studies will be reviewed and
the apparent consensus approach for test methods discussed. Currently, the
state-of the-art is in its infancy, and the development of standard methods is
not feasible; however, it is crucial that there be an understanding of the
significant effect which these methods have on sediment quality evaluations. It
is anticipated that recommended methods and this guide will be routinely updated
to reflect progress in our understanding of sediments and how to best study them.
1.2 There are several regulatory guidance documents (1-16) concerned
with sediment collection and characterization procedures, which might be
important for individuals performing Federal or State agency-related work.
Discussion of some of the principles and current thoughts on these approaches can
be found in Dickson et al., 1987 (17).
1 Biological Sciences Dept., Wright State University, Dayton, OH 45435
2 Great Lakes Environmental Research Lab, 2205 Commonwealth BlvcL, Ann Arbor, MI 48105
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1.3 This guide is arranged as follows:
Section
Scope 1
Referenced Documents 2
Terminology 3
Summary of Guide 4
Significance and Use 5
Interferences 6
Apparatus 7
Safety Hazards 8
Sampling and Transport 9
Storage 10
Collection of Interstitial Water 11
Characterization 12
Manipulation 13
Quality Assurance 14
Report 15
References
1.4 Field collected sediments might contain potentially
toxic materials and thus should be treated with caution to
minimize occupational exposure to workers. Worker safety must
also be considered when working with spiked sediments containing
various organic or inorganic contaminants, or both; and those
that are radio-labeled. Careful consideration should be given to
those chemicals which might biodegrade, volatilize, oxidize, or
photolyze during the test period.
1.5 This standard does not purport to address all of the
safety problems associated with its use. It is the
responsibility of the user of this standard to establish
appropriate safety and health practices and determine the
applicability of regulatory limitations prior to use. Specific
hazard statements are given in Section 8.
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Referenced Documents
2.1 ASTM Standards:
D 1129 Definitions of Terms Relating to Water
0 4387 Guide for Selecting Grab Sampling Devices for
Collecting Benthic Macroinvertebrates
D 4822 Guide for Selection of Methods of Particle
Size Analysis of Fluvial Sediments (Manual
Methods).
D 4823 Guide for Core Sampling Submerged,
Unconsolidated Sediments
E 380 Practice for Using the International System
of Units (SI) (the Modernized Metric System)
E 729 Practice for Conducting Acute Toxicity Tests
with Fishes, Macroinvertebrates, and
Amphibians
E 943 Definitions of Terms Relating to Biological
Effects and Environmental Fate
E 1023 Guide for Assessing the Hazard of a Material
to Aquatic Organisms and Their Uses
E 1367 Guide for Conducting Solid Phase 10-day
Static Sediment Toxicity Tests with Marine
and Estuarine Amphipods
E ??? Guide for Conducting Solid Phase Sediment
Toxicity Tests with Freshwater Invertebrates
E 1295 Guide for Conducting Three Brood Renewal
Toxicity Tests with Ceriodaphnia dubia
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5
3. Terminology
3.1 The words "must", "should", "may", "can", and "might"
have very specific meanings in this guide. "Must" is used to
express an absolute requirement, that is, to state that the test
ought to be designed to satisfy the specified condition, unless
the purpose of the test requires a different design. "Must" is
only used in connection with the factors that directly relate to
the acceptability of the test. "Should" is used to state that
the specified condition is recommended and ought to be met in
most tests. Although a violation of one "should" is rarely a
serious matter, violation of several will often render the
results questionable. Terms such as "is desirable", "is often
desirable", and "might be desirable" are used in connection with
less important factors. "May" is used to mean "is (are) allowed
to", "can" is used to mean "is (are) able to", and "might" is
used to mean "could possibly". Thus, the classic distinction
between "may" and "can" is preserved, and "might" is never used
as a synonym for either "may" or "can".
3.2 Definitions. For definitions of terms used in this
guide, refer to Guide E 729, Definitions E 943, and Definitions
D 1129, and Guide D 4387; for an explanation of units and
symbols, refer to Practice E 380.
4. Summary of Guide
4.1 This guide provides a review of widely used methods to
collect, store, characterize, and manipulate sediments for
toxicity testing. Where the science permits, recommendations are
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6
provided on which procedures are appropriate, while identifying
their limitations.
5. significance and Us*
5.1 Sediment toxicity evaluations are a critical component
of environmental quality and ecosystem impact assessments, used
to meet a variety of research and regulatory objectives.
5.2 The manner in which the sediments are collected,
stored, characterized, and manipulated can greatly influence the
results of any sediment quality or process evaluation.
Addressing these variables in a systematic and uniform manner
will aid interpretations of sediment toxicity or bioaccumulation
results and may allow comparisons between studies.
6. Interferences
6.1 Maintaining the integrity of a sediment environment during
its removal, transport, and testing in the laboratory is extremely
difficult. The sediment environment is composed of a myriad of
microenvironments, redox gradients, and other interacting
physicochemical and biological processes. Many of these
characteristics influence sediment toxicity and bioavailability to
benthic and planktonic organisms, microbial degradation, and chemical
sorption. Any disruption of this environment complicates
interpretations of treatment effects, causative factors, and in situ
comparisons. For additional information see Section 9.
7. Apparatus
7.1 A variety of sampling, characterization, and manipulation
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7
methods exist using different equipment. These are reviewed in
Sections 9 through 14.
7.2 Cleaning; Test chambers and equipment used to prepare
and store dilution water and stock solutions should be cleaned
before use. New and used sample containers should be washed
following these steps: (1) non-phosphate detergent wash, (2)
triple water rinse, (3) water-miscible organic solvent wash,
(acetone followed by pesticide grade hexane (2,8)), (4) water
rinse, (5) acid wash (such as 5% concentrated hydrochloric
acid), and (6) triple rinse with deionized-distilled water.
Altering this cleaning procedure might result in problems. Many
organic solvents might leave a film that is insoluble in water
(Step 3). A dichromate-sulfuric acid cleaning solution can
generally be used in place of both the organic solvent and the
acid (Steps 3 through 5), but it might attack silicone adhesive.
(See 9.10 for cleaning during sample collection.)
8. Safety Hazards
8.1 Many substances can adversely affect humans if adequate
precautions are not taken. Information on toxicity to humans
(18) and recommended handling procedures of toxicants (19) should
be studied before tests are begun with any contaminant or
sediment. Health and safety precautions should be considered
before beginning a test.
8.2 Field collected sediments might contain a mixture of
hazardous contaminants and/or disease causing organisms such that
proper handling to avoid human exposure is important. Therefore,
skin contact with all test materials and solutions should be
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8
minimized by such means as wearing appropriate protective gloves
especially when washing equipment or putting hands in dilution
water over sediments, or into sediments. Proper handling
procedures might include: 1) sieving and distributing sediments
under a ventilated hood or an enclosed glove box, 2) enclosing
and ventilating the toxicity test water bath, and 3) using
respirators, aprons, safety glasses, and gloves when handling
potentially hazardous sediments. Special procedures might be
necessary with radiolabeled test materials (20) and with
materials that are, or are suspected of being, carcinogenic (19).
8.3 Disposal of sediments, dilution water over sediments,
and test organisms containing hazardous compounds might pose
special problems. For tests involving spiking sediments with
known toxicants, removal or degradation of the toxicant(s) before
disposal is sometimes desirable. Disposal of all hazardous
wastes should adhere to the requirements and regulations of the
Resource Conservation and Recovery Act and any relevant State or
local regulations.
9. sampling and Transport
9.1 Sediments have been collected for a variety of
chemical, physical, toxicological and biological investigations.
These collections have been made with both a series of grab
sampling devices and core samplers (See Table 2, Guide D 4823).
The advantages and disadvantages of the various collection
methods have been previously reported (3,4) and are summarized in
Table 1. All sampling methods disturb the sediment integrity to
a degree. For purposes of sediment toxicity evaluations it is
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9
important to obtain sediments with as little disruption as
possible, to allow for realistic laboratory evaluations of in
situ conditions. Choosing the most appropriate sediment sampler
for a study will depend on the sediments characteristics, the
efficiency required, and the study objectives. Several
references are available which discuss the various collection
devices (3,4,21,22,23). The efficiency of these samplers for
benthic collections have been compared and in general the grab
samplers are less efficient collectors than the corers but are
easier to handle, work in heavier seas, often require fewer
personnel and are more easily obtained (21,23-31).
9.2 The principal disadvantage of dredge samplers varies;
common problems are shallow depth of penetration and presence of
a shock wave that results in loss of the fine surface sediments.
Murray and Murray (32), however, described a dredge usable in
heavy seas which quantitatively samples the top 1 cm of sediment
and retains fine materials. Other grab samplers that
quantitatively sample surface sediments have been described by
Grizzle (33). The depth profile of the sample may be lost in the
removal of the sample from the sampler. Dredge sampling promotes
loss of not only fine sediments, but also water soluble compounds
and volatile organic compounds present in the sediment.
9.3 Studies of macroinvertebrate sampling efficiency with
various grab samplers have provided useful information for
sampling in sediment toxicity and sediment quality evaluations.
The Ekman dredge is the most commonly used sampler for benthic
investigations (21). The Ekman*s efficiency is limited to less
compacted, fine-grained sediments, as are the corer samplers.
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10
The most commonly used corer is the Kajak-Brinkhurst corer. In
more resistant sediments the Petersen, PONAR, and Smith-Mclntyre
dredges are used most often (21). Based on studies of benthic
macroinvertebrate populations, the sediment corers are the most
accurate samplers, followed by the Ekman dredge, in most cases
(21). For resistant sediments, the PONAR dredge was the most
accurate and the Petersen the least (21). A comparison of
sampler precision showed the van Veen sampler to be the least
precise; the most precise were the corers and Ekman dredge (21).
9.4 Many of the problems associated with dredge samplers
are largely overcome with the corers. The best corers for most
sediment studies are hand-held polytetrarfluoroethylene plastic,
high density polyethylene, or glass corers (liners), or large
box-corers. The corers can maintain the integrity of the
sediment surface while collecting a sufficient depth.
Furthermore, the box core can be sub-cored or sectioned at
specific depth intervals, as required by the study. The box
corer, unfortunately, is large and cumbersome; thus, it is
difficult to use. Other coring devices which have been
successfully used include the percussion corer (34) and vibratory
corers (35-37).
9.5 Corer samplers also have several limitations. Most
corers do not work well in sandy sediments; dredge samplers or
diver-collected material remain the only current alternatives.
In general, corers collect less sediment than dredge samplers
which may provide inadequate quantities for some studies. Small
cores tend to increase bow waves (that is, disturbance of surface
sediments) and compaction, thus altering the vertical profile.
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11
However, these corers provide better confidence limits and
spatial information when multiple cores are obtained (21,24,38-
41). As shown by Rutledge and Fleeger (42) and others, care must
be taken in subsampling from core samples, since surface
sediments might be disrupted in even hand-held core collection.
They recommend subsampling in situ or homogenizing core sections
before subsampling.
9.6 Studies of sediment toxicity, interstitial waters,
microbiological processes, or chemical fate probably will require
core sampling to best maintain the complex integrity of the
sediment. When obtaining cores from shallow waters one must
ensure that the vessel does not disturb the sediments prior to
sampling (30). Most of the studies in the literature employed
grab samplers although box corers (43-45), gravity corers (46)
and hand collection (47-49) methods are reported with increasing
frequency. For additional information of various core types see
reference USEPA (4).
9.7 Subsampling, compositing, or homogenization of
sediment samples is often necessary and the optimal methods will
depend on the study objectives. Important considerations
include: loss of sediment integrity and depth profile; changes
in chemical speciation via oxidation and reduction or other
chemical interactions; chemical equilibrium disruption resulting
in volatilization, sorption, or desorption; changes in biological
activity; completeness of mixing; and sampling container
contamination. In most studies of sediment toxicity, it is
advantageous to subsample the inner core area (not contacting the
sampler) since this area is most likely to have maintained its
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12
integrity and depth profile and not be contaminated by the
sampler. Subsamples from the depositional layer of concern, for
example, the top 1 or 2 cm should be collected with a nonreactive
sampling tool, such as, a polytetrafluoroethylene lined
calibration scoop (50). Samples are frequently of a mixed depth
but a 2 cm sample (51) is the most common depth obtained,
although depths up to 40 ft have been used in some dredging
studies. For some studies it is advantageous or necessary to
composite or mix single sediment samples (16,50). Composites
usually consist of three to five grab samples. Subsamples are
collected with a nonreactive sampling scoop and placed in a
nonreactive bowl or pan. The composite sample should be stirred
until texture and color appear uniform.
9.8 Due to the large volume of sediment which is often
needed for toxicity or bioaccumulation tests and chemical
analyses, it might not be possible to use subsampled cores
because of sample size limitations. In those situations, the
investigator should be aware of the above considerations and
their possible affect on test results as they relate to in situ
conditions.
9.9 Assessment of in situ sediment toxicity or
bioaccumulation is aided by collection and testing of reference
and control samples. For purposes of this guide, a reference
sediment is defined as a sediment possessing similar
characteristics to the test sediment but without anthropogenic
contaminants. Sediment characteristics, such as particle size
distribution and percent organic carbon, should bracket that of
the test sediment. If there is a wide range of test sediment
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13
types, the reference sediment characteristics should be in an
intermediate range unless the test species is affected by
particle size. The appropriate ASTM guides for marine and
freshwater invertebrates should then be consulted to determine
the particle size requirements of the test species. It is
preferable that reference sediments be collected from the same
aquatic system, located close to, and have similar physical,
chemical, and biological characteristics to the test sediment.
In some situations, the reference sediment might be toxic due to
naturally occurring chemical, physical, or biological properties.
For this reason, it is important to also test the toxicity of
control sediments. The reference sediment test results might be
analyzed as either a treatment or as a control variable,
depending on the study objectives. For purposes of this guide, a
control sediment might consist of natural or artificially
prepared sediments of known composition and of consistent quality
that have been used in prior sediment toxicity tests or
culturing, and for which baseline data exists which shows they do
not cause toxicity. Control sediments have been successfully
used in toxicity evaluations (52).
9.10 When collecting sediment grab samples, it is
important to clean the sampling device, scoop, spatula, and
mixing bowls between sample sites. The cleaning procedure can
follow that outlined in Section 7 or the following (53): 1) soap
and water wash, 2) distilled water rinse, 3) methanol rinse, 4)
methylene chloride rinse, and 5) site water rinse. Waste
solvents should be collected in labelled hazardous waste
containers.
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14
9.11 In most cases the transport conditions for the
samples were not specified in the references reviewed. Where
conditions were specified, the sediments were usually transported
whole, in both plastic, polyethylene (54-56), and glass
(48,49,57) containers and transported under refrigeration or on
ice (48,49,51,57-62).
9.12 Collection, transport, storage, and test chamber
material composition should be chosen based on a consideration of
sorption effects, sample composition, and contact time. For
example, in sediments where organics are of concern, brown
borosilicate glass containers with Polytetrafluoroethylene (PTF)
lid liners are optimal, while plastic containers are recommended
for metal samples. PTF or high density polyethylene containers
are relatively inert and optimal for samples contaminated with
multiple chemical types. Additionally, polycarbonate containers
have been shown not to sorb metal species (63). Additional
information on sample containers, preservation, storage times and
volume requirements, in regards to chemical analyses, are
available in other guidance documents (3-6,10,16). In many cases
these criteria are applicable to toxicity test chamber
requirements.
10. storage
10.1 Containers for storage were generally not specified
although it was assumed that the containers were the same as the
transport containers, where specified, and were generally
polyethylene (see 9.12). Where sediments contain volatile
compounds, transport and storage should be in air tight PTF or
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15
glass containers with PTF-lined screw caps. For further
information on storage requirements for chemical analyses see
Table 2.
10.2 Drying, freezing, and cold storage conditions all
affect toxicity (17,64-69). Often the storage time of sediments
used in toxicity tests was not specified and where specified
ranged from a few days (70) to one year (55). Storage of
sediments after arrival at the laboratory was generally by
refrigeration at 4 C (54-56,58-62,67,70-73). Significant changes
in metal toxicity to cladocerans and microbial activity have been
observed in stored sediments (68,74). Recommended limits for
storage of metal-spiked sediments have ranged from within 2 days
(64) to 5 days (70) to 7 days (75,76). A study of sediments
contaminated with nonpolar organics found that interstitial water
storage time did not affect toxicity to polychaetes when samples
were frozen (77). Cadmium toxicity in sediments has been shown
to be related to acid volatile sulfide (AVS) complexation (78) .
When anoxic sediments were exposed to air, AVS were rapidly
volatilized. AVS is apparently the reactive solid phase sulfide
pool that binds metal, thus reducing toxicity. If a study
objective is to investigate metal toxicity and the sediment
environment is anoxic, then exposure to air might reduce or
increase toxicity due to oxidation and precipitation of the metal
species or loss of acid volatile sulfide complexation. It is
generally agreed that sediments to be used for toxicity testing
should not be frozen (17,67,69,70,75,79).
10.3 Although risking changes in sediment composition,
several studies elected to freeze samples (51,67,80-84). Fast-
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16
freezing of sediment cores has been recommended for chemical
analyses; however, this alters sediment structure and profile
distortion occurs (42). Freezing has been reported to inhibit
oxidation of reduced iron and manganese compounds (81). It has
also been recommended for stored sediments which are to be
analyzed for organics and nutrients (85).
10.4 Interstitial water chemistry changed significantly
after 24 h storage (86,87), even when stored at in situ
temperatures (87). Coagulation and precipitation of the humic
material was noted when interstitial water was stored at 4 C for
more than one week (88). Oxidation of reduced arsenic species in
pore water of stored sediments was unaffected for up to 6 weeks
when samples where acidified and kept near 0 C, without
deoxygenation. When samples were not acidified, deoxygenation
was necessary (89).
10.5 In summary* sediments for toxicity tests and chemical
analyses are typically refrigerated or placed on ice in
polyethylene containers during transport. If, in addition,
samples are to be used for chemical analyses, then the
appropriate container should be used as described above. The
storage conditions should be refrigeration at 4 C and under
anoxic conditions if appropriate (10,16,90). It has been shown
that sediments can be stored at 4 C for up to 12 months without
significant alterations in toxicity (91). Limits to storage time
before testing, therefore, appear to be a function of both
sediment and contaminant characteristics. While it is prudent to
complete the testing of sediments with a minimum of storage time
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17
(probably less than 2 weeks) this may not be possible for any
number of reasons.
11. Collection of Interstitial Water
11.1 Isolation of sediment interstitial water can be
accomplished by several methods: centrifugation, squeezing,
suction, and equilibrium dialysis. In general, methods for recovery
of relatively large volumes of interstitial water from sediments are
limited to either centrifugation (57,88,92,93) or squeezing (94-97).
Other methods, such as suction (98), gas pressurization (50), in
situ samplers (99), and equilibration by using dialysis membrane or
a fritted glass sampler (100-103), do not produce large quantities
of interstitial water. In the case of the dialysis, sufficient time
must be allowed to ensure that the sample has come to equilibrium
with the interstitial water. The suction and dialysis equilibrium
methods are most useful for laboratory studies. Some pore water
constituents, for example, dissolved organic carbon or
dimethylsulfide, might significantly affected by the collection
method (99). Other constituents, such as, salinity, dissolved
inorganic carbon, ammonia, sulfide, and sulfate, might not be
affected by collection methods providing oxidation is prevented
(99). If sediments are anoxic, all steps involved in sample
processing might need to be conducted in inert atmospheres to
prevent oxidation of reduced species (99,104,105).
11.2 If interstitial water is collected by centrifugation and
filtration, then effects on the interstitial chemistry need to be
considered after centrifugation. Centrifugation followed by 2/xm
filtration yielded similar metal concentrations to dialysis methods
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18
(106). However, filtration with glass fiber or plastic filters is
not appropriate in some cases and has been shown to remove nonpolar
organics (107). Centrifugation at 7600 x g with glass contact only
was shown to be superior to filtration methods (107). Other studies
have produced contrary results, recommending filtration with
polycarbonate filters (98,108). Filtration is normally conducted to
remove particles with a 0.45 Mm pore size, however 0.20 /un or
smaller pore size membranes have been recommended (81). Removal of
all bacteria and colloidal materials might require filter pore sizes
of less than 0.2 jm. Immediate collection of interstitial water is
recommended since chemical changes might occur even when sediments
are stored for short periods at in situ temperatures (87) (see
10.4) .
12. Characterization
12.1 The characteristics that have been most often measured in
sediments are moisture content, organic carbon or volatile matter
content, and particle size. When attempting to characterize a
sediment, quality assurance should always be addressed (3,4,16).
Sediments, by their nature, are very heterogenous; they exhibit
significant temporal and spatial heterogeneity in the laboratory and
in situ. Replicate samples should be analyzed to determine the
variance in sediment characteristics and analytical methods.
Sediment characterization will depend on the study objectives and
the contaminants of concern, however, a minimum set of
characteristics should be included which are known to influence
toxicity and will aid data interpretation: in situ temperature,
particle size distribution, moisture or interstitial water content,
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19
ash free weight, organic carbon (determined by titration or
combustion), pH, Eh, acid volatile sulfides, ammonia, and cation
exchange capacity. Many of the methods of characterization have
been based on analytical techniques for soils and waters and the
literature should be consulted for further information
(15,23,109,110).
12.2 The moisture content of sediments is measured by drying
the sediments at 50 to 105*C to a constant weight (23).
12.3 Volatile matter content is often measured instead of, and
in some cases in addition to, organic carbon content as a measure of
the total amount of organic matter in a sample. This measurement is
made by ashing the sediments at high temperature and reporting the
percent ash free dry weight (7,111,112). Although the exact method
for ashing the sample is often not specified, the normally accepted
temperature is 550 ± 50 C (16,23).
12.4 Carbon fractions which may be of importance in
determining toxicant fate and bioavailability include: total
organic carbon (16,113-115), dissolved organic carbon (88),
dissolved inorganic carbon, sediment carbonates, and reactive
particulate carbon (116,117). Reactive particulate carbon is that
portion which equilibrates with the aqueous phase. The organic
carbon content of sediments has been measured by wet oxidation which
is also useful for the determination of the organic carbon content
of water (118). Organic carbon analyses have also been conducted by
titration (119), modification of the titration method (120), or
combustion after removal of carbonate by the addition of HC1 and
subsequent drying (73).
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20
12.5 Particle sizing of sediments can be measured by numerous
methods (15,121, see Guide D 4822) dependent on the particle
properties of the sample (122). Particle size distribution is often
determined by wet sieving (2,15,16,23,123). Particle size classes
might also be determined by the hydrometer method (124,125), the
pipet method (15,126), settling techniques (127), X-ray absorption
(123,126) and laser light scattering (128). The pipet method may be
superior to the hydrometer method (129). To obtain definite
particle sizes for the fine material, a Coulter (particle size)
counter method might be employed (130,131). This method gives the
fraction of particles with an apparent spherical diameter. Another
potential method for determining the particle size distribution of a
very fine fraction is through the use of electron microscopy (132).
The collection technique for the very fine materials can result in
aggregation to larger colloidal structures (132-135). Comparisons
of particle sizing methods have shown that some produce similar
results and others do not. These differences might be attributed to
differences in the particle property being measured, that is, the
Malvern Laser Sizer and Electrozone Particle Counter are sizing
techniques, and the hydrophotometer and SediGraph determine
sedimentation diameter based on particle settling (122,136-138). It
is preferable to use a method which incorporates particle settling
as a measure, as opposed to strictly sediment sizing.
12.6 Various methods have been recommended to determine
bioavailable fractions of metals in sediments (78,139-141). One
extraction procedure, cation exchange capacity, provides information
relevant to metal bioavailability studies (109). Amorphic oxides of
iron and manganese, and reactive particulate carbon have been
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2:
implicated as the primary influences on metal sorption potential in
sediments (81,140,142-144). Measurement of acid volatile sulfide
(AVS) and divalent metal concentrations associated with AVS
extraction provides insight into metals availability in anaerobic
sediments (78). Easily extractable fractions are usually removed
with cation displacing solutions, for example, neutral ammonium
acetate, chloride, sodium acetate, or nitrate salts (145).
Extraction of saltwater or calcareous sediments, however, is often
complicated by complexation effects or dissolution of other sediment
components (141,146). Other extractants and associated advantages
and disadvantages have been recently discussed (141,144,147,148).
Some extractants which have been successfully used in evaluations of
trace metals in nondetrital fractions of sediments are EDTA or HC1
(141,149,150). Metal partitioning in sediments might be determined
by using sequential extraction procedures which fractionate the
sediments into several components such as interstitial water, ion
exchangeable, easily reducible organic and residual sediment
components (93,148,151,152). Unfortunately at this time no one
method is clearly superior to the others (147). This might be due,
in part, to site specific characteristics which influence
bioavailability, for example, desorption and equilibration
processes.
12.7 pH is important for many chemicals and can be measured
directly (23) or in a 1 to 1 mixture of sediment/soil to water
(153).
12.8 Eh measures are particularly important for metal
speciation and for determining the extent of sediment oxidation.
Redox gradients in sediments often change rapidly over a small depth
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22
and are easily disturbed. Care must be taken in probe insertion to
allow equilibration to occur when measuring Eh. These measurements
are potentiometric and measured with a platinum electrode relative
to a standard hydrogen electrode (23).
12.9 Biochemical oxygen demand and chemical oxygen demand
might provide useful information in some cases (23). Sediment
oxygen demand might also be a useful descriptor; however, a wide
variety of methods exist (90,154-157).
12.10 Analysis of toxicants in sediments is generally
performed by standard methods such as those of the EPA (2,23).
Soxhlet extraction is generally best for organics but depends on
extraction parameters (158,159). Concentrations are generally
reported on a dry weight basis.
13. Manipulation
13.1 Manipulation of sediments is often required to yield
consistent material for toxicity testing and laboratory experiments.
The manipulations reviewed in this section are: spiking (dosing)
regimes for laboratory and control sediments; mixing; sieving for
attainment of maximal particle sizes; dilutions for concentration-
effect determinations; elutriates; capping; air drying; and
sterilization. For discussion of subsampling, compositing, or
homogenization effects see 9.7.
13.2 Spiking — The spiking method to be used is contingent on
the study objectives. For example, when attempting to mimic in situ
conditions, sediment cores should be spiked by adding aqueous or
suspended sediment solution of toxicants to the overlying water
column; or when investigating dredging effects or conditions of
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2:
sediment perturbation where toxicant sorption processes are
accelerated, mixing toxicants into sediment slurries may be
advantageous. When investigating the source of sediment toxicity 01
interactive effects of sediment toxicants, it is useful to spike
both reference and control sediments with the toxicant of concern
present in the test sediment. Mixing time should be limited to a
few hours and temperatures kept to a minimum, due to the rapid
alterations which occur in the sediment's physicochemical and
microbiological characteristics, which thereby alter bioavailability
and toxicity. Recalcitrant organics and some metals, for example,
cadmium and copper, might be mixed for extended periods without
adverse effects (see 9 through 12 for additional discussion).
13.3 Organic compounds are generally added via a carrier
solvent such as acetone or methanol to ensure that they are soluble
and that they remain in solution during mixing. While organic
compounds are generally added in an organic carrier, metals are
generally in aqueous solutions. Compounds are also added to water
overlying sediments and the compound allowed to sorb with no mixing
(71,160-167). Occasionally the carrier has been added directly to
sediment (52,82-84,112,137,168-171) and the carrier evaporated
before addition of water. This approach does not seem to result in
compounds being sorbed to sediment at the same sites as dosing under
aqueous conditions (172). Word et al. (107) compared several
sediment-labelling techniques using methylene chloride, ethanol, and
glycine as carriers. They found glycine was superior when mixed
with sediment for 7 days. In most cases, the compound is either
coated on the walls of the flask and an aqueous slurry (sediment and
water in various proportions) added, or the carrier containing
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24
mixture is added directly to the slurry. When the sediment to water
ratio is adjusted for optimal mixing, sediments that are too dense
to mix by slurrying in water have been successfully mixed using a
rolling mill (72). Other mixing techniques may be used for spiking
specific sediments but care should be taken to ensure complete
mixing and analyses of spiked compounds run to ensure that labelling
is uniform in the mixed material. The use of a polar, water soluble
carrier such as methanol has little effect on the partitioning of
nonpolar compounds to dissolved organic matter at concentrations up
to 15% carrier by volume (173). Another study, however, shows that
changes in partitioning of a factor of approximately two, might well
occur with 10 % methanol as a cosolvent for anthracene sorption
(174). Thus, caution should be taken to minimize the amount of
carrier used. The time between the spiking of the compounds and the
use of the test sediment has been variable (46,47,70,72,73,80,111,
168,175) and does seem to effect the biological availability of
compounds (37,67,175).
13.4 Highly volatile compounds have been spiked into sediments
in a similar manner to the less volatile materials using cosolvents
and mixing in an aqueous slurry by shaking. These experiments were
tested immediately in covered flow through systems (108).
13.5 If a solvent other than water is used, both a sediment
solvent control and a sediment negative control or reference
sediment; or both, must be included in the test. The solvent
control must contain the highest concentration of solvent present
and must use solvent from the same batch used to make the stock
solution (see Practice E 729).
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25
13.6 Because the organic carbon content of the sediments might
be one of the most important characteristics affecting the
biological availability of contaminants, modifications of the carbon
content have been made in many studies. Methods used include
dilution with clean sand (55,56,62,108); although humics (170) and
other organics such as sheep manure (52) have also been added. Such
dilutions also change the particle composition and the size
distribution of the particles; thus, results from such experiments
should be interpreted with care. The organic carbon content has
also been altered by the use of combustion (14,52). Combustion may
alter the type of carbon as well as oxidize some of the inorganic
components thus altering greatly the characteristics of the
sediment.
13.7 A variety of methods have been used to spike sediments
with metals. The two principal categories of methods are 1) metal
addition directly to the sediment which is mixed and then water
added (64,68,176-178), or 2) addition of the metal to the overlying
waters (80,166,179,180). Thorough mixing of spiked sediments has
been accomplished using the rolling mill tecnigue, Eberbach and gyn
rotary shakers.
13.8 Equilibration and mixing conditions vary widely in
spiking studies. The duration of contact between the toxicant and
sediment particles can affect both the partitioning and
bioavailability of the toxicant. This effect apparently occurs
because of an initial rapid labile sorption followed by movement of
the toxicant into resistant sorption sites or in the particle (181-
183). Because of the kinetically controlled changes in the
partitioning that results in changes in bioavailability (175,184,
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26
185), the contact tine can be important when spiking sediments.
Bounds on the sorption time can be estimated from the partition
coefficient for the sediment following the calculations in
Karickhoff and Morris (182). In addition, it is important to
recognize that the quantity of toxicant spiked might exceed the
complexation capacity of the test sediment system and not allow
reactions to attain equilibrium. These phenomenon will complicate
test result interpretation (68,147).
13.9 Mixing and sieving are two other manipulations of
sediments that are often performed before toxicity testing
(46,52,58-60,67,70,72,111,112,163,168,170,175,186). Sediment
samples have been sieved for a variety of reasons including the
removal of large debris and stones thereby increasing the samples
homogeneity and method replicability; the increased ease of counting
organisms; the increased sediment handling and subsampling; the
ability to study influence of particle size on toxicity,
bioavailability, or contaminant partitioning. Sieving of material
to a specific size fraction might alter the concentration of
contaminant in the sediment by removing large, low sorptive
materials.
13.10 Toxicants and organic carbon concentrations tend to be
higher with fine grained sediments (that is, clay and silt) due to
increased surface area (in relation to the weight of the sample) and
sorptive capacity. Measuring size fractions of less than 63 /im has
been recommended in contaminant studies, particularly for metals
(172,187). In studies of sediment metal concentrations, normalizing
to the less than 63 jum size fraction was superior for describing
metal binding in sediments, as compared to sediment concentrations
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27
normalized to dry weight, by organic carbon content, or corrected by
a centrifugation procedure (172). Small size fractions are
characteristic of depositional areas in aquatic systems; however,
sieving of sediments from non-depositional sites to obtain ' fine
fraction might significantly alter the sediment characterist .s.
The usual sieve size for toxicity testing is greater than 500 /Ltm.
If sieving is performed it should be done for all samples to be
tested including control and reference sediments.
13.11 Mixing of various layers of sediments might result in
either dilution or enhancement of concentrations. The sediment
quality will be influenced by the depth of sampling, depth of
biological activity, contaminant solubility and partitioning
characteristics, and depth of the contaminant concentration peak
which is dependent on historical contamination and sedimentation
rates for the study site, see Section 10 for additional relevant
discussion.
13.12 Another manipulation of sediments for toxicity testing
is sediment dilution. In order to obtain concentration-effect
information in solid phase sediment toxicity evaluations, differing
concentrations of the test sediment should be used. Currently,
there is little information available on the most appropriate method
for diluting test sediments to obtain a graded contaminant
concentration or concerning the methodological effects of s n a
dilution. A "clean" noncontaminated sediment should be used ns the
"diluent" which optimally consists of physicochenr —«1
characteristics similar to the test sediment, such as organic
matter/carbon, particle size, but does not contain elevated (above
background) levels of the toxicants of concern. Refer to the
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28
preceding sections for relevant information.
13.13 Many studies of sediment toxicity have been conducted on
the elutriate or water-extractable phase (188). This method was
developed to assess the effects of dredging operations on water
quality. Sediments are shaken in site or reconstituted water (1 to
4 volume to volume ratio) for 30 min. The water phase is then
separated from the sediment by centrifugation, followed by
filtration of the supernatant through a 0.45 jum filter when
conducting some tests, such as algal growth assays. The filtration
step may be removed depending on the study objectives (see Section
11 for interferences).
13.14 Sediment pollution remediation alternatives might
include capping the contaminated sediments with "clean" sediments.
Laboratory design of such experiments should vary the depth of both
the contaminated sediments and the capping sediment layers to
evaluate contaminant transport via physicochemical and biological
(bioturbation) processes.
13.15 Sometimes sediments have been air dried before use
(56,168,189,190) but these sediments have generally been used for
laboratory studies after some additional manipulation, such as
spiking sediments with various levels of contaminants for
concentration-effect data (111,190). Air drying would result in
losses of volatile compounds and might result in changes in the
sediment characteristics, particularly particle size (see Section
10). The presence of air and air drying have all been shown to
change metal availability and complexation (141).
13.16 Sterilization of sediments to inhibit biological
activity has been performed in some studies. Autoclaving is used in
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29
most cases (191). Other sterilization techniques have included:
antibiotic addition, addition of chemical inhibitors such as HgCl or
sodium azide, or gamma irradiation. The technique chosen should be
contingent on study objectives. Antibiotics, such as streptomycin
and ampicillin, have been successfully used in sediment studies
(192,193). Some antibiotics, however, are labile and light
sensitive, or readily bind to organic matter. Mercuric chloride
appears to be superior to sodium azide as a bacteriocide.
Autoclaving is the least desirable method as it causes the greatest
alteration to the sediments physical and chemical characteristics.
In studies requiring sterility, it is crucial that a sterility
control be incorporated.
14. Quality Assurance
14.1 Quality assurance guidelines (3,4,10,16) should be
followed. Quality assurance considerations for sediment modeling,
QA-QC plans, statistical analyses (for example, sample number and
location) and sample handling have been addressed in-depth (10).
14.2 Sediment heterogeneity significantly influences studies
of sediment quality, contaminant distribution, and both benthic
invertebrate and microbial community effects. Spatial heterogeneity
might result from numerous biological, chemical, and physical
factors and should be considered both horizontally (such as, the
sediment surface) and vertically (that is, depth). Accumulation
areas with similar particle size distributions might yield
significantly different toxicity patterns when subsampled (79,194);
therefore, an adequate number of replicates should be processed to
determine site variance. When determining site variance one should
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30
consider within sample (that is, subsample) variance, analytical
variance (for example, chemical or toxicological), and the sampling
instruments' accuracy and precision. After these considerations a
sampling design can be constructed which addresses resource
limitations and study objectives.
14.3 As stated in previous sections, the methodological
approach used, such as, number of samples, will be dependent on the
study objectives and sample characteristics. For information on
sediment heterogeneity, splitting, compositing, controls, or
determining sample numbers, sampler accuracy and precision, and
resource requirements, there are a number of references available
(4,10,21,85,172,195,196).
15. Report
15.1 Documentation: The record of sediment collection,
storage, handling, and manipulation should include the following
information either directly or by reference to existing documents.
Published reports should contain enough information to
clearly identify the methodology used and the quality of the results.
15.1.1 Name of test and investigator(s), name and location of
laboratory, and dates of starting and ending of sampling and
sediment manipulation;
15.1.2 Source of control, reference or test sediment, method
for handling, storage and disposal of sediment;
15.1.3 Source of water, its chemical characteristics, and a
description of any pretreatment;
15.1.4 Methods used for, and results (with confidence limits)
of, physical and chemical analyses of sediment; and
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31
15.1.5 Anything unusual about the study, any deviation from
these procedures, manipulations, and any other relevant information.
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TABLE 1 (continued). Summary of Bottom Sampling Equipment
33
Jill.
Advantaget
Disadvantage*
PONAR Grab Sampler
Deep lakes, rivers, and estuaries.
Useful on sand, silt, or clay.
Host universal grab sampler. Adequate
on most substrates. Large sample
obtained Intact, permitting subsamp ling.
Shock wave from descent may
disturb "fines". Possible
Incomplete closure of jaws
results In sample loss.
Possible contamination from
metal frame construction.
Sample must be further prepared
for analysis.
BNH-53 Piston Corer
Waters of 4-6 feet deep when
used with extension rod. Soft
to semi-consolidated deposits.
Piston provides for greater
sample retention.
Cores must be extruded on
site to other containers -
Metal barrels introduce risk of
metal contamination.
Van Veen
Deep lakes, rivers, and estuaries.
Useful on sand, silt, or clay.
Adequate on most substrates. Large
sample obtained intact, permitting
subsampling.
Shock wave from descent may
disturb "fines". Possible
incomplete closure of jaws
results In sample loss.
Possible contamination from
metal frame construction.
Sample must be further prepared
for analysis.
Mffl-60
Sampling moving waters from •
fixed platform
Streamlined configuration allows
sampling where other devices could not
achieve proper orientation.
Possible contamination from
metal construction. Sub-
sampling difficult. Not
effective for sampling fine
sediments.
Petersen Grab Sampler
Deep lakes, rivers, and estuaries.
Useful on most substrates.
Large sample; can penetrate
most substrates.
Heavy, may require winch.
No cover lid to permit sub-
sampling. All other
disadvantages of Ekman and
Ponar.
Shipek Grab Sampler
Used primarily In marine waters
and large Inland lakes and
reservoirs.
Sample bucket may be opened to
permit subsampling. Retains fine
grained sediments effectively.
Possible contamination from
metal construction. Heavy,
may require winch.
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34
TABLE 1 (continued). Suimary of Bottom Sampling Equipment
Device USS Advantage* pjsadvantafles
Orange-Peel Grab Deep lake*, river*, and estuaries. Designed for sampling hard substrates. Loss of fines. Heavy - may
SMith-Mclntyre Grab Useful on most substrates. requires winch. Possible
Metal contamination.
Scoops, Drag Buckets Various environments depending Inexpensive, easy to handle. Loss of fines on retrieval
on depth and substrate. through water column.
~ ~ " (modified, 193)
1 Comments represent subjective evaluations.
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35
TABLE 2.
Contaminant
Sampling Containers, Preservation Requirements, and Holding Time* for Sediment Samples*
(EPA, 196,197). See also Rochon and Chevalier (160).
Acidity
Alkalinity
Ammonia
Sulfate
Sulfide
Sulfite
Nitrate
Nitrate-Nitrite
Nitrite
Oil and Grease
Organic Carbon
Metals
Chromium VI
Mercury
Metals except above
Organic ConDounds
Extractables (including
phthalates, atrosamines
organochlorine pesticides
PCS'* artroaromatics,
isophorone, Polynuclear
aromatic hydrocarbons.
haloethers, chlorinated
hydrocarbons and TCDD)
Extractables (phenols)
P.G
P.G
P.G
P.G
P.G
P.G
P.G
P.G
P.G
G
P,G
P.G
P.G
P.G
G, t
1
G, t
Container* Preservation
Cool, 4C
Cool, 4C
Cool, 4C
Cool, 4C
Cool. 4C
Cool, 4C
Cool, 4C
Cool, 4C
Cool, 4C
Cool, 4C
Cool, 4C
Cool, 4C
teflon-lined cap Cool, 4C
Purgables (halocarbons
and aromatic*)
Purgables (acrolein and
acrylonitrate)
Orthophosphate
Pesticides
Phenols
Phosphorus (elemental)
Phosphorus, total
Chlorinated organic
compounds
teflon-lined cap Cool, 4C
G, teflon-lined Cool, 4C
septum
G, teflon-lined Cool, 4C
septua
P.G Cool, 4C
G, teflon-lined cap Cool, 4C
P.G Cool, 4C
G Cool, 4C
P,G Cool, 4C
G, teflon-lined cap Cool, 4C
Holding Time
14 days
14 days
28 days
28 days
28 days
48 h
48 h
28 days
48 h
28 days
28 days
40 h
8 days
6 months
7 days (until extraction)
30 days (after extraction)
7 days (until extraction)
30 days (after extraction)
14 days
3 days
48 h
7 days (until extraction)
30 days (after extraction)
28 days
48 h
28 days
7 days (until extraction)
30 days (after extraction)
Taken from EPA 600-4-84-075 and EPA 600-4-85-048, see also Ref. 85.
Polyethylene (P) or Glass (G)
-------
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168. Foster, G. D., Baksi, S. M., and Means, J. C., "Bioaccumulation
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may attend. If you feel that your comments have not received a
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19103."
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Elutriate Preparation
FIELD-COLLECTED SEDIMENT ELUTRIATE PREPARATION
I. General:
This SOP describes the procedures for homogenizing stored sediment samples and preparation of
sediment elutriate samples for toxitity testing. Sediment is often a storage reservoir for many
contaminants introduced into surface waters. These contaminants may include polychlorinated
biphenyls, polymiclear aromatic compounds and inorganic contaminants including heavy metals.
Contaminants present in sediment may include carcinogens, mutagens, or potentially toxic
compounds. Toxirity tests will be started before chemical analyses can be completed in most cases.
Since field sediments may contain potentially toxic materials they should be treated with caution to
minimize occupational exposure to workers.
H. Safety.
A. Personal precautions.
1. Workers must always be aware of possible points of contamination as described by
the supervisor. Hands should always be kept away from the eyes and mouth. After
completion of a manipulation involving sediment or the removal of possibly
contaminated laboratory clothing (gloves, lab coat, etc.), the hands, forearms, and
other areas of suspected contact should be washed with hand soap and water at a
sink located within the laboratory work area. Do not use organic solvents to clean
the skin. These solvents may increase penetration of the contaminant into the skin.
B. Containment devices.
1. All work with sediment will be performed in an appropriate containment device.
Procedures involving sediment will not be conducted on an open bench due to the
potential hazard of generating contaminated dusts, aerosols, or fumes. Hoods,
glove boxes, and enclosed vented water baths for testing and rooms equipped with
once pass ventilation are used to minimize the worker exposure to contaminants
associated with sediment. All containment devices will be constructed out of
smooth, unbreakable material, such as Teflon", stainless steel, polyethylene,
fiberglass, or plexiglass.
C. Work surfaces.
1. All work surfaces potentially exposed to sediments must be covered with Teflon0
sheets, plastic trays, dry absorbent plastic-backed paper, foil, or other impervious or
disposable material. If a surface becomes contaminated or if a spill occurs, the
work surface should be decontaminated or disposed of immediately.
III. Storage of sediment
A. Solid-phase sediment and sediment elutriates and extracts.
1. Solid-phase sediment and sediment elutriates and extracts will be stored at 4° C in
air-tight containers b the dark.
a. All samples must be accompanied with proper identification and sample
tracking information and can be temporarily stored at 4°C in refrigerators
located in the laboratory work areas.
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IV. Homogenization of sediments
A. Sediment homogenization or manipulation increases the chances for occupational exposure.
During sediment homogenization or other manipulations, the number of investigators in the
laboratory work area should be minimized. Other workers in the building must be notified
of the handling of the sediment.
B. All mixing of solid-phase sediment or preparation of sediment extracts or elutriates wi- be
performed either in a fume hood or while wearing the appropriate clothing and respiratory
protective equipment. If the containers holding sediment are removed from the hood, an
intermediate non-breakable container must be used.
V. Elutriate preparation
A. Required equipment.
1. Balance capable of weighing at least 1500 ± .01 grams.
2. Polypropylene centrifuge bottles.
3. Modified 60 cc polypropylene disposable syringes.
a. Remove tip from syringe barrel.
b. Drill a 3/8 inch opening at end of barrel.
c. Wash plunger and barrel in soap and water, rinse with well water, rinse
with 10% HC1 followed by 3 D.I. water rinses.
4. Elutriate mixing apparatus.
a. The elutriate mixing apparatus consists of a 1/10 HP, 14 rpm, shaded pole
gear motor (Dayton Model 3M136A) supported horizontally by a metal
frame constructed of 1 inch square tubing. The motor drive is attached via
a flexible bushing to the end of a stainless steel box measuring 31 x 23 x 18
cm. The top of the box is removable and secured to the box with two
wing-nuts. The interior of the box is divided into 6 compartments
measuring 10 x 10 x 18 cm. Each compartment accepts one 1000 mL
polypropylene centrifuge bottle. The motor rotated the stainless steel box
end over end on two lubricated pillow block bearing assemblies.
5. Large volume centrifuge.
a. The centrifuge is an International Equipment Company Model PR-7000,
refrigerated, large capacity centrifuge equipped with the Model 966 rotor.
Maximum Relative Centrifugal Force with the 966 rotor is 7400 x G at 6900
rpm.
B. Temperature of manipulations.
1. Sediment samples for elutriate preparation will be taken immediately after
homogenization. All manipulations will be done at room temperature (~20°C)
except for centrifugation which will be performed at ~4°C.
C. Method.
1. Preparing elutriates with 1000 mL centrifuge bottles.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Elutriate Preparation
a. Individually weigh 10 - 1000 mL. centrifuge bottles and caps to be used in
sample preparation and obtain a mean weight.
b. Round the mean weight obtained up to the nearest gram and record this
weight (e.g. if the mean weight of 10 bottles is 90.89 grams, round this
value to 91 grams).
c. Place a dean centrifuge bottle (without the cap) on balance. Tare bottle to
0.00 grams.
d. Transfer 200.00 ± .05 grams of sediment using a modified 60 cc
polypropylene disposable syringe to the centrifuge bottle.
e. Remove the centrifuge bottle containing the weighed sediment from the
and re-zero the balance.
f. Replace the centrifuge bottle (with sediment) and cap on the pan and add
dilution water until the combined weight of the bottle, sediment, cap and
bottle equals 1000 grams plus the rounded average gram weight of the
containers obtained in step V-C-l-b (above). For example if the average
rounded weight of the centrifuge bottles was 91.0 grams, water would be
added to the centrifuge bottle containing 200 grams of sediment until the
combined weight of the bottle, cap, sediment and water was 1091.00 ± .05
grams.
2. Preparing elutriates with 250 mL centrifuge bottles.
a. Individually weigh 10 - 250 mL. centrifuge bottles and caps to be used in
sample preparation and obtain a mean weight.
b. Round the mean weight obtained up to the nearest gram, and record this
weight (e.g. if the mean weight of 10 bottles is 34.56 grams, round this
value to 35 grams).
c Place a dean centrifuge bottle (without the cap) on balance.
d. Transfer 50.00 ± .05 grams sediment using a modified 60 cc polypropylene
disposable syringe to the centrifuge bottle.
e. Remove the centrifuge bottle containing the weighed sediment from the
pan and re-zero the balance.
f. Replace the centrifuge bottle (with sediment) and cap on the pan and add
dilution water until the combined weight of the bottle, sediment, cap and
bottle equals 250 grams plus the rounded average gram weight of the
containers obtained in step V-C-2-b above. For example if the average
rounded weight of the centrifuge bottles was 35.0 grams, water would be
added to the centrifuge bottle containing 50 grams of sediment until the
combined weight of the bottle, cap, sediment and water was 285.00 ± .05
grams.
3. Mixing elutriates.
a. Centrifuge bottles containing the appropriate weights of water and
sediment are placed in the elutriate mixing apparatus and rotated end over
end for 30 minutes at 12 rpm per minute.
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b. After samples have mixed for 30 minutes, re-weigh all centrifuge bottles to
0.01 grains prior to transferring them to the centrifuge. Record weights.
All bottles must be within ± 0.20 grams of each other. If necessary, add
sufficient SJVDP water with a pipet to bottles containing weights below this
range.
FAILURE TO BRING ALL BOTTLES WITHIN ± 0.2 GRAMS PRIOR TO
CENTRIFUGATION MAY RESULT IN ROTOR IMBALANCE AND
DAMAGE TO THE CENTRIFUGE.
4. Centrifuging elutriates.
a. Transfer bottles to centrifuge buckets. Position of bottles in the rotor is
not important if all bottles are within the ± 0.02 gram range.
(1) Bottles must be centrifuged in pairs and placed in opposite buckets
in the rotor. If an odd number of bottles are to be centrifuged,
prepare a blank bottle that weighs within ± 0.20 grams of the
opposite bottle.
b. Samples are centrifuged at 5,000 rpm (7000 x G) for 15 min. at 4 °C.
(1) Check that temperature displayed is 4 ± 2 °C.
(2) Set SPEED thumb-wheel switch to 18.5 min. (adding 3.5 minutes
to the 15 min. centrifuge time allows centrifuge to attain the set
speed).
(3) Set BRAKE thumb-wheel switch to 2.
(4) Set ACCELERATION thumb-wheel switch to 1.
(5) Press Start/Stop button.
5. Removing elutriate from centrifuge bottles.
a. The overlying water from each centrifuge bottle containing sediment from
the same site is poured through a clean 50 mesh stainless steel standard
sieve into a dean 3.0 L glass bottle and mixed.
b. Sub-samples of the elutriate are obtained from the 3.0 1. glass bottle and
are stored in appropriate containers in the dark at 4°C.
6. Elutriate sub-sampling and analyses.
a. Chemical characterization of the elutriate sample may include the
following: pH, total water hardness, alkalinity, conductivity, ammonia,
dissolved oxygen, and turbidity.
b. A 500 mL sample of the elutriate will be placed in 500 mL teflon-lined
bottle for metal analysis. These samples will be acidified to pH 1.7-2.0
using Baker instant analyzed acid. About 0-5 mL of acid in 500 mL of
elutriate sample should achieve this range in pH. The sample will be
stored at 4°C until analysis for metals.
c. The elutriate sample may need to be filtered before using in toxitity testing
(e.g.. Selenastrum capricornut^nn or Ames testing).
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop
ASTM Designing Sediment Tests
Draft #1
10/19/90
This document is in process of development and is for ASTM
committee use only. It shall not be reproduced or circulated or
quoted, in whole or in part, outside of ASTM committee activities
except with the approval of the chairman of the committee having
jurisdiction or the President ofthe Society.
GUIDE FOR DESIGNING BIOLOGICAL TESTS WITH SEDIMENTS
Jim Dwyer1, Bill Goodfellow2, Chris IngersolT
Anne Keller3, Wayne McCulloch2, Skip Missimer4
Dick Peddicord2, Charles Pittinger5, and John Scott8
1.0 Scope
1.1 As contamination of freshwater, estuarine, and marine ecosystems continues to be reduced through
the implementation of regulations governing both point and non-point source discharges, there is a growing
emphasis and concern regarding historical inputs and their influence on water and sediment quality. Many
locations hi urban areas exhibit significant sediment contamination which poses a continual and long-term threat
to the health of benthic communities and other species inhabiting these areas (NOAA, 1988). Benthic
communities are an important component of many food chains leading to humans and it is becoming increasingly
important to identify contaminated sites to properly manage remediation and resource use.
12 Biological tests with sediments are an efficient means for evaluating sediment contamination because
they provide information complementary to chemical characterizations and ecological surveys (Chapman, 1988).
Acute sediment toxicity tests can be used as screening tools in the early phase of an assessment hierarchy that
ultimately could include chemical measurements or bioaccumulation and chronic effects tests. Sediment tests
have been applied in both marine and freshwater environments (Swartz 1987; Chapman, 1988; Lamberson and
Swartz, 1988). Sediment tests have been used for dredge material permitting, site ranking for remediation,
1 National Fisheries Contaminant Research Center, 4200 New Haven Road, Columbia, MO 65201
2 EA Engineering Science and Technology, INC., Hunt Valley/Loveton Center, 15 Loveton Circle,
Sparks, MD 21152
3 St. Johns Water Management District, P.O. Box 1429, Palatka, FL 32178-1429
4 PH. Glatfelter Company, 228 South Main St., Spring Grove, PA 17362
5 The Proctor and Gamble Company, Ivorydale Technical Center - IN24, Cincinnati, OH 45217
6 Science Applications International Corporation, U.S. EPA-ERL, South Ferry Road, Narragansett, RI
02882
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recovery studies following management actions, and trend monitoring. A particularly important application is in
establishing contaminant-specific effects and the processes controlling contaminant bioavailability.
2.0 Application
2.1 This document provides general interpretative guidance on the selection, application and
interpretation of biological tests with sediments. As such, it serves as a preface to other ASTM documents
describing: methods for sediment collection, storage and manipulation (ASTM E 1391); toxicity tests with marine
(ASTM E 1367) and freshwater organisms (ASTM E 1383); and bioaccumulation studies. This guide serves as
an introduction and summary of sediment testing; it is not meant, however, to provide specific guidance on test
methods. Rather, its intent is to provide information necessary to:
2.1.1 Select a sediment exposure strategy that is appropriate to the assessment need of the toxicity
test. For example, a suspended phase exposure is relevant to evaluation of dredged sediments
for disposal at a dispersive aquatic site.
2.1.2 Select the test organism and biological endpoints that are appropriate to the desired exposure
and aquatic resources at risk. For example, the potential for water quality problems and
subsequent effects on oyster beds may dictate the use of sediment elutriate exposures with
bivalve larvae.
o Establish an experimental design consistent with the objectives of the sediment evaluation. The
use of appropriate controls is particularly important here.
o Determine which statistical procedures should be applied to the analysis of the data, and define
the limits of applicability of the resultant analyses in the data interpretation.
3.0 Organization (To be drafted)
4.0 Hazard statement/Safety precautions
4.1 Many substances may pose health risks to humans if adequate precautions are not taken.
Information on toxicity to humans, recommended handling procedures, and chemical and physical properties of
the test material should be studied before a test is begun and made aware to all personnel involved (6,7,8).
Contact with test materials, overlying water and sediments should be minimized.
4.2 Many materials can adversely affect humans if precautions are inadequate. Skin contact with
test materials and solutions should be minimized by such means as wearing appropriate protective gloves,
laboratory coats, aprons, and safety glasses, and by using dip nets, sieves or tubes to remove test organisms from
overlying water. When handling potentially hazardous sediments the proper handling procedures may include (a)
sieving and distributing sediments under a ventilated hood or in an enclosed glove box, (b) enclosing and
ventilating the water bath, and (c) use of respirators, aprons, safety glasses, and gloves. Field collected sediments
may contain potentially toxic materials and should be treated with caution to minimi?/-, occupational exposure to
workers. Worker safety should also be considered when working with spiked sediments containing various
organic or inorganic compounds, compounds that are radiolabeled, and with materials that are, or are suspected
of being, carcinogenic or teratogenic (7).
43 Careful consideration should be given to those chemicals which might biodegrade, biotransform
to more toxic components, volatilize, combust, oxidize, or photolyze during the test period.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM Designing Sediment Tests
4.4 Health and safety precautions and applicable regulations for disposal of stock solutions, test
organisms, sediments, and overlying water should be considered before beginning a test (ASTM Standard D
4447).
5.0 Applicable Documents
5.1 ASTM Documents
E 380 Standard for Metric Practice
D 1129 Definitions of Terms Relating to Water
E 1023 Guide for Assessing the Hazard of a Material to Aquatic Organisms and Their Uses
E 943 Standard Definitions of Terms Relating to Biological Effects and Environmental Fate
E 1367 Guide for Conducting Solid Phase 10-Day Static Sediment Toxicity Tests with Marine and
Estuarine Infaunal Amphipods
E 1391 Guide for Collection, Storage, Characterization, and Manipulation of Sediments for
Toxicological Testing.
E 1383 Guide for Conducting Sediment Toxicity Tests with Freshwater Invertebrates
6.0 Terminology
6.1 The words "must", "should", "may", "can", and "might" have very specific meanings in this guidance.
"Must" is used to express an absolute requirement, that is, to state that the test ought to be designed to satisfy a
specific condition, unless the purpose of the test requires a different design. "Must" is only used in connection
with the factors that apply directly to the acceptability of the test. "Should" is used to state that the specified
conditions are recommended and ought to be met in most tests. Although a violation of one "should" is rarely a
serious matter, violation of several will often render the results questionable. Terms such as "is desirable", "is
often desirable", and "might be desirable" are used in connection with less important factors. "May" is used to
mean "is (are) allowed to", "can" is used to mean "is (are) able to", and "might" is used to mean "could possibly".
Thus, the classic distinction between "may" and "can" is preserved, and "might" is never used as a synonym of
either "may" or "can".
62 sediment - is used to denote a naturally occurring paniculate material which has been transported
and deposited at the bottom of a body of water. The term can also be applied to an artificially prepared
substrate within which the test organisms can interact.
6.2.1 whole sediment - is distinguished from elutriate, and resuspended sediments, in that the whole,
intact sediment is used to expose the organisms, not a form or derivative of the sediment.
6.2J2 dean -- denotes a sediment (or water) that does not contain concentrations of test materials or
xenobiotics which cause apparent stress to the test organisms or reduce then* survival.
63 elutriate - refers to the water or solvent used to elute contaminants from the sediment and is then
used in aquatic exposures.
6.4 suspended -- is a slurry of sediment and water used to expose the organisms.
6.5 overlying water ~ the water placed over the solid-phase of a sediment in the test chamber for the
conduct of the biological test, and may also include the water used to manipulate the sediments.
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6.6 interstitial water — the water within a wet sediment that surrounds the sediment particles, expressed
as the percent ratio of the weight of the water in the sediment to the weight of the wet sediment.
6.7 spiking — the experimental addition of a test material such as a chemical or mixture of chemicals,
sewage sludge, oil, particulate matter, or highly contaminated sediment to a clean negative control or reference
sediment, such that the toxicity of the material added can be determined.
6.8 concentration — the weight or volume of test material(s) associated with a weight or volume of test
sample.
6.9 exposure -- is contact with a chemical or physical agent.
6.10 toxicity - is the property of a material or combination of materials to adversely affect organisms.
6.11 bioaccumulation - the net uptake of a material by an organism from its environment through direct
exposure or ingestion.
6.12 Control sediment - a sediment essentially free of contaminants (USEPA-COE 1990). Any
contaminants in control sediment originates from the global spread of pollutants and does not reflect any
substantial input from local or non-point sources (Lee et al. 1989). The comparison of the test sediment to the
control sediment is a measure of any toxicity from the test sediment beyond inevitable background contamination
(Lee et al. 1989). The control sediment is used to assess the acceptability of the test and provide evidence of the
health and quality of the test animals (Nelson et al. 1990).
6.13 Reference sediment - a sediment substantially free of contaminants (USEPA-COE 1990). The
reference sediment may be used as an indicator of localized sediment conditions exclusive of the specific pollutant
input of concern. Such sediment would be collected near the site of concern and would represent the background
conditions resulting from any localized pollutant inputs as well as the global input (Lee et al. 1989). This is the
manner in which reference sediment is used in the dredged material evaluations (EPA-COE 1990).
6.14 For definitions of other terms used in this practice, refer to Standards E 729, E 943, D 1129, E
1023, and E 1241. For an explanation of units and symbols, refer to Standard E 380.
7.0 Summary of Guide
7.1 This guide provides general guidance and objectives for conducting biological tests with sediments.
Detailed technical information on the conduct and evaluation of specific sediment tests is included in other
documents referenced in this guide.
7.2 Neither this guide nor any specific test methodology can adequately address the multitude of
technical factors that must be considered when designing and conducting a specific investigation. Therefore, the
intended use of this document is not to provide detailed guidance but rather to assist the investigator in
developing technically sound and environmentally relevant biological tests that adequately address the questions
being posed by a specific investigation.
8.0 Sediment Test Rationale (Significance and Use)
8.1 Contaminated sediments may have adverse effects on natural populations of aquatic organisms.
Sediment dwelling organisms may be directly exposed to contaminants by the ingestion of sediments and by the
uptake of sediment-associated contaminants from interstitial and overlying water. Contaminated sediments may
directly affect water column species by serving as a source of contaminants to overlying waters or a sink for
contaminants from overlying waters. Organisms may also be affected when contaminated sediments are
suspended in the water column by natural or human activities. Water column species and non-aquatic species
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM Designing Sediment Tests
may also be indirectly affected by contaminated sediments by the transfer of contaminants through aquatic-
terrestrial food chains.
8.2 The test methodologies described herein may be used and adapted for incorporation in basic and
applied research projects to further clarify the ecological effects of contaminated sediments. These same
methods may also be used in the development and implementation of regulatory programs designed to prevent
the contamination of sediments and manage sediments that are already contaminated.
83 Sediment tests with aquatic organisms can be used to quantify the acute and chronic toxicity and the
bioavailability of new and presently used materials. In many cases, consideration of the adverse effects of
sediment-associated contaminants is only one part of a complete hazard assessment of manufactured compounds
that are intentionally released to the environment (e.g., pesticides, herbicides) and those released only
inadvertently through the manufacturing process (e.g., through wastewater effluents).
8.4 Sediment tests can be used to develop dose-response relationships for individual toxicants by spiking
clean sediments with varying concentradons of a test chemical and determining the concentration that elicits the
target response in the test organism. In a similar fashion, sediment tests can be designed to determine the effects
that die physical and chemical properties of sediments have on die bioavailability and toxicity of compounds.
85 Properly designed and conducted sediment tests can provide valuable information needed to make
decisions regarding die management of contaminated sediments from hazardous waste sites and other
contaminated areas. Biological tests with sediments can also be used to make defensible management decisions
on die dredging and disposal of potentially contaminated sediments from rivers and harbors.
9.0 Sediment Test Types
9.1 Recent reviews have summarized methods for assessing die toxicity of marine [2^3] and freshwater
[4,5] sediments to benthic organisms. Those methods are provided in Table 1 and Table 2, for marine and
freshwater tests, respectively.
92 The selection of a specific toxicity test type is intimately related to die objectives of die sediment
evaluation program. These assessments, whedier they be for monitoring, regulatory, or research purposes, should
be guided by a set of null hypotheses which define die appropriate exposure route and die endpoint of interest.
93 Organism exposure methods most commonly employ die whole sediment in die bedded phase, but
suspended and elutriate phase exposures have also been used More recendy, methods have been developed to
test pore waters directly and to prepare organic extracts for testing. The relationship between toxicity resulting
from these latter exposures and what may be found in situ, however, is not well defined.
9.4 Programs seeking to characterize or rank sediments on a basin-wide or regional scale typically use
whole sediment, solid phase exposures. Regulatory or permitting programs for dredged material disposal at a
containment site should also evaluate tiiis exposure route. Disposal at a dispersive site, or concerns over
resuspension and transport of in-place sediments, would suggest use of suspended phase or elutriate exposures.
95 Mediods have been developed to isolate and test die toxicity of elutriates (e.g., USEPA-COE 1977)
or sediment interstitial water (e.g., Ankley et al. 1990) to aquatic organisms. The elutriate test was developed for
assessing die potential acute effects of open-water disposal of dredged material Tests witil elutriate samples are
used to estimate die water soluble constituents dial may be released from sediment to die water column during
-------
disposal operations (Shuba et al. 1977). Toxicity tests of the elutriate with water column organisms have
generally indicated little toxitity is associated with the discharge material (Lamberson and Swartz 1988).
However, elutriates have been reportedly more toxic than interstitial water samples (Giesy and Hoke 1989).
9J.I For many benthic invertebrates, the toxicity and bioaccumulation of sediment-associated
contaminants such as metals, and non-ionic organic contaminants are correlated with the concentration of these
chemicals in the interstitial water (Ankley et al. 1990 ammonia). The sediment interstitial water toxicity test was
developed for assessing the potential jn .sjQi effects of contaminated sediment on aquatic organisms. Once the
interstitial water (or elutriate) has been isolated from the whole sediment, the toxicity testing procedures would
be similar to effluent toxicity testing with non-benthic species. If benthic species are used as test animals, they
may be stressed by the absence of sediment (Lamberson and Swartz 1988).
952 Examination of organic extracts may have specific uses when whole sediments have a
predetermined toxicity and cause-effect relationship. However, caution must be exercised in the use of organic
extracts because the resultant contaminant interactions within a sediment matrix have not been determined.
9.6 Biological responses in sediment toxicity tests range from genotoxic effects to individual organism
responses to alterations in community levels of organization. Because of its ease of interpretation, the response
criterion that is most commonly employed has been lethality. This endpoint is generally insensitive to sediment
contaminants unless appropriately sensitive species, such as amphipods, are used. The application of sublethal
toxicity tests has been limited because of the uncertainty in relating these responses to ecologically relevant
endpoints such as survival and population dynamics. Behavioral responses of infaunal organisms, such as
emergence from the sediments are indicative of potential ecological effects because the animals may be subject to
predation. Many biochemical and genetic endpoints, e.g., enzyme induction and chromosome aberration, are
indicative of exposure to specific classes of chemicals, and are useful from that perspective. Sublethal tests which
show the most promise are those using growth and reproduction as response parameters. These are relevant
endpoints that can be used as predictors of potential population effects. Most of these tests, however, are still in
development and are limited in their application. Tests combining lethality with growth and reproduction have
been developed and routinely applied using freshwater and marine organisms.
9.7 The selection of the proper response parameter can also be predicated on the goals of the evaluation
program, but the choice is often based on available resources, tune, and test methods. Sublethal endpoints are
generally the preferred responses, but they are more difficult to interpret and/or the data are more costly to
generate. Sediment screening programs commonly use simple reliable testes, e.g. amphipod mortality, bacterial
biolummcsccncc or sea urchin fertilization. The latter two tests are conducted on either pore waters or organic
extracts. In depth evaluations of single sediments, as in U.S. Army Corps of Engineers dredging evaluations, are
more likely (o involve a more complex suite of tests including life cycle scale responses or long-term
bioaccum illation studies. Specific sublethal responses such as genotoxicity or enzyme induction may be used to
identify contaminant-specific exposures.
10.0 Test Organisms
10.1 Once the exposure routes and endpoints of interest have been established, there are several criteria
that need to be considered when selecting the appropriate test species (Shuba 1981, Swartz 1987). Ideally, the
test species should:
o have a lexicological (sediment) database demonstrating sensitivity to a range of toxicants,
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM Designing Sediment Tests
o be readily available through field collection or culture,
o be easily maintained in the laboratory,
o be ecologically or economically important,
o have a broad geographical distribution,
o be indigenous to the site being evaluated or closely related to an inhabitant,
o be tolerant to a broad range of sediment geochemical characteristics (e.g., organic carbon and
grain size), and
o be capatible with selected exposures and endpoints.
Of these criteria, demonstrated sensitivity to contaminants, ecological relevance, and tolerance to varying
sediment geochemical characteristics are the most important. The use of indigenous species that are ecologically
important and easily collected is often very straightforward, however, many indigenous species at a contaminated
site may be insensitive to contaminants. These might present a greater concern relative to their bioaccumulation
potential. With the exception of some amphipods, few test organisms have broad sediment or water toxicity
databases. Additionally, many organisms can be maintained in the laboratory long enough for acclimation to test
conditions, but very few are easily cultured. Widespread toxicity testing will require cultured organisms or the
use of standard source populations which can be transported without experiencing excessive mortality.
10.2 Sensitivity is related to the degree of contact between the sediment and the organism. Feeding
habits including the type of food and feeding rate will control the dose of contaminant from sediment (Adams
1987). Infaunal deposit-feeding organisms can receive a dose of sediment contaminants from three sources:
interstitial water, whole sediment, and overlying water. Benthic invertebrates may selectively consume particles
with higher organic carbon and higher contaminant concentrations. Organisms hi direct contact with sediment
may also accumulate contaminants by direct adsorpdon to the body wall or exoskeleton, or by absorption through
the integument (Knezovich et al. 1987). Thus, estimates of bioavailability will be more complex for epibenthic
animals that inhabit both the sediment and the water column. Some benthic organism are exposed primarily by
detrital feeding (Boese 1988 sab). Detrital feeders may not receive most of their body burden directly from
interstitial water. For certain higher Kow compounds, uptake by the gut can exceed uptake across the gill
(Landrum 1989, Boese et al. 1990). However, for many benthic invertebrates, the toxicity and bioaccumulation of
sediment-associated contaminants such as metals, kepone, fluorathene, and organochlorines are highly correlated
with the concentration of these chemicals in the interstitial water (Ankley et al. 1990).
103 The marine tests cover a broad spectrum of taxa and feeding types including crustaceans, bivalves,
polychaetes and fish. Tests using amphipods have received a great deal of attention because field surveys have
shown them to be absent from contaminated sites. This sensitivity has led to the development of routine methods
using the burrowing amphipod Rhepoxynius abronius. This ten day acute toxicity test has recently been adapted
for use with other amphipod species and has been established as a standard method by ASTM. Since 1977, the
U.S. Army Corps of Engineers dredging permit program has routinely required tests with three species: a bivalve,
a polychaete and a fish or shrimp, incorporating organisms which burrow into the sediment and those inhabiting
the water column. Broad applications of these protocols reveal that these tests are not as sensitive as those with
amphipods and the latter recently have been recommended for permit programs.
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10.4 Sediment tests in freshwater utilize a number of different species. Whole sediment tests with the
amphipod Hvalella azteca generally start with juvenile animals and are conducted for up to four weeks until
reproductive maturation (Nelson et al. 1990). Although a direct measurement of amphipod reproduction is
appealing, the quantitative isolation of young amphipods from sediment is difficult because of their small size
(<2mm). Indirect measures of reproduction, such as time to reproductive maturation, or the number of eggs or
young carried in the marsupium are more easily quantified than the number of young produced. Moreover, the
total number of young produced during the exposure may reflect not only a direct effect on reproduction, but
may also be affected by a reduction in adult survival (Ingersoll and Nelson 1990).
Tests with Chironomus tentus are generally started with 2nd instar larvae (10-14 d old) and continued for
10 to 17 d until the 4th instar; larval survival or growth is the measure of toxicity (Nelson et al. 1990). Exposures
of C. tentans that started with 1st instar larvae or that measured adult emergence have met with only limited
success [39, Nebeker et aL 1988 BECT). Whole sediment testing procedures with C. riparius are started with 1
to 3 day old larvae and continued through pupation and adult emergence (Nelson et al. 1990). Midge exposures
started with older larvae may underestimate midge sensitivity to toxicants. For instance, 1st instar £. tentans
larvae were 6 to 27 times more sensitive than 4th instar larvae to acute copper exposure [34,39], and 1st instar C.
riparius larvae were 127 times more sensitive than 2nd instar larvae to acute cadmium exposure [44].
Sediment toxicity tests with mayflies and dadocerans are generally conducted for up to 10 days (Bahnick
et al. 1981, Nebeker et al. 1984, Giesy et aL 1990 ETC 9:2). Survival and molting frequency are the toxicity
endpoints monitored in the mayfly tests and survival, growth, and reproduction are monitored in the cladoceran
tests. While dadocerans are not in direct contact with the sediment, they are frequently in contact with the
sediment surface and are likely exposed to both water soluble and paniculate bound contaminants in the
overlying water and surface sediment (Stemmer and Burton 1990 ASTM). dadocerans are also one of the more
sensitive groups of organisms used in toxicity testing.
The most frequently described sediment test methods for oligochaetes are acute toxicity testing
procedures (e.g., Keilty et al. 1988 AECT 17:95-101). Wiederholm et al. (1987) describe methods for conducting
up to 500 day oligochaete exposures with growth and reproduction as the toxicity endpoint. Recently, Reynoldson
et al. (in prep.) describe a 28 d test started with sexually mature Tubifex tubifex. In this shorter test, effects on
growth and reproduction can be monitored and the duration of the exposure makes the test more useful for
routine sediment toxicity assessments with oligochaetes. Oligochaetes have complex life cydes and reproductive
strategies and as a result laboratory culturing requirements have prohibited their use in toxicity testing (Dillion
and Gibson 1985).
105 Because of the database that has been developed with existing tests, it is recommended that, for
whole sediment exposures, either phoxocephalid or ampeliscid amphipods be used in marine tests. For
freshwater applications, hyalellid amphipods, midge larvae, or mayfly larvae would be appropriate. As new
methods are developed, it will be important to establish each method's sensitivity relative to a benchmark
procedure for comparative purposes (Chapman 1988). The marine benchmark should be the Rhepoxvnius
abronius ten day acute test and the freshwater benchmark should be the Hvalella azteca. Although sublethal
tests with whole sediments are rare, aggressive attempts should be made to develop tests using growth and
reproduction endpoints with marine and freshwater amphipods.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM Designing Sediment Tests
10.6 Multispecies/microcosm tests can also be used to evaluate potential ecosystem responses to
contaminated sediments. However, results from multispecies or microcosm tests are more difficult to interpret
due to interactions and limited reference literature (13, Prater and Hoke 1980).
11.0 Experimental design considerations
11.1 Sample methods
11.1.1 Purpose of the study—the probable source and type of contamination and the objectives of the
study, should be evaluated before developing the sampling regime. The number of samples taken and method of
sampling may vary depending on the objectives of the study (8,13,11,1).
11.12. The number of replicate samples taken at a site should be determined based on a preliminary
survey of sediment variability at the site. The mean and standard deviation of the replicates can be used to
calculate a minimum number of replicates (13,1).
11.13 In general, both toxitity and bioaccumulation tests require at least two exposures - a control and
one or more test treatments. The experimental unit for each test is the exposure chamber. Typically a sediment
sample is split into four or more test chambers. Individual observations obtained from within an individual
chamber should not be used as replicate observations. Replicate chambers for a particular sediment provide an
estimate of the variability within the test system and are not sediment sample replicates.
11.1.4 There are several acceptable methods of sampling sediments, e.g. corers and grabs or dredges.
Grabs or dredges (e.g., Ponar or Eckman) are appropriate when sediments are known to be unstratified with
respect to the contaminants of concern. If the contaminants are in strata or if their accumulation rates are of
interest, one of several core samplers should be used. Pb210 or Cst37 dating can be performed on cores to
identify the thickness of the mixed layer (13). See ASTM 1391 for additional details.
11.2 Sample handling and preservation are discussed in ASTM E 1391 and depend on the type of
chemical characterization that will be performed. The use of clean sampling devices and sample containers is
essential to ensure the accurate determination of sediment contamination (13,1).
113 Physical and chemical characterization of sediments may include loss on ignition, percent water,
grain size, total organic carbon, total phosphorus, nitrogen forms, trace metals and organic compounds, pH, total
volatile solids, biological oxygen demand, chemical oxygen demand, cation exchange capacity, Eh, pE, total
inorganic carbon, acid volatile sulfides, and ammonia (8,11,1).
11.4 Overlying Water ~ Besides being available in adequate supply, overlying water used hi toxicity tests,
and water used to hold organisms before testing, should be acceptable to test species and uniform in quality. To
be acceptable to the test species, the water must allow satisfactory survival and growth, without showing signs of
disease or apparent stress, such as discoloration, or unusual behavior.
11.4.1 Natural overlying water should be uncontaminated and of constant quality and should meet the
specifications established hi ASTM E 729. Water should be characterized in accordance with ASTM E 729 at
least twice each year, and more often if (a) such measurements have not been determined semiannually for at
least two years, or (b) if surface water is used.
11.42 A natural overlying water is considered to be of uniform quality if the monthly ranges of the
hardness, alkalinity, and specific conductance are less than 10% of their respective averages and if the monthly
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range of pH is less than 0.4 units. Natural overlying waters should be obtained from an uncontaminated well or
spring, if possible, or from a surface water source. If surface water is used, the intake should be positioned to
minimize, fluctuations in quality and the possibility of contamination and maximize the concentration of dissolved
oxygen and to help ensure low concentrations of sulfide and iron. Chlorinated water should not used for, or in
the preparation of, overlying water because residual chlorine and chlorine-produce ..xidants are toxic to many
aquatic animals and dechlorination is often incomplete.
11.43 For certain applications the experimental design might require use of water from the test
sediment collection site.
11.4.4 Reconstituted water is prepared by adding specified amounts of reagent grade chemicals to high
quality distilled or deionized water (see ASTM E 729). Acceptable water can be prepared using deionization,
distillation, or reverse-osmosis units. Conductivity, pH, hardness and alkalinity should be measured on each
batch of reconstituted water. If the water is prepared from a surface water, total organic carbon or chemical
oxygen demand should be measured on each batch. Filtration through sand, rock, bag, or depth-type cartridge
filters may be used to keep the concentration of paniculate matter acceptably low. The reconstituted water
should be intensively aerated before use, except that buffered soft fresh waters should be aerated before, but not
after, addition of buffers. Problems have been encountered with some species in some fresh reconstituted waters,
but these problems can be overcome by the aging the reconstituted water for one or more weeks.
H-5 Test Design
11.5.1 Materials used to construct test chambers may include glass, stainless steel, silicone and plastics
that have been properly prepared and tested for toxicity (ASTM E 1367, E 1383).
11.5.2 The use of site water or reconstituted water in toxicity tests may depend on the type of test to be
performed and the time lapse between sample collection and test initiation. 11.5.3 Static sediment
toxicity tests are the simplest to perform and have been commonly used. In such tests, water overlying the
sediment is not changed during the test period, but may be added to replace that which has evaporated. Since
changes in water quality may affect the availability of contaminants to the test organisms, static exposures are
more appropriate for acute tests (7-10 days).
11-5.4 Flow-through exposure chambers are suggested for use in chronic tests or with larger animals.
Since water is renewed on a continual basis, fewer water quality changes are likely due to the buildup of waste
products or interactions between the sediment and overlying water.
11.5.5 General water quality (variables such as pH, dissolved oxygen, ammonia, and temperature) in the
test chambers should meet culture and maintenance requirements for the test organisms. These parameters
should be monitored and recorded on a frequency appropriate to the test length. For example, if the test
duration is only a few days, daily monitoring should be performed. However, if the test will continue for weeks
or months, measurements may be reduced to every other day or every few days.
11.5.6 The depth of sediment in test chambers may vary depending on the organism being tested, its size
and degree of burrowing activity, and its sediment processing rate. The latter should be determined prior to the
beginning of a sediment toxicity test (13).
115.7 Control and/or reference sediments should be used in each sediment test. A standard reference
sediment is a well characterized sediment containing a known amount of a specific pollutant (13) which may be
prepared by spiking in the laboratory with an appropriate compound, e.g. organic or metal compound. The
10
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM Designing Sediment Tests
standard reference sediment is useful as an indicator of test organism variability among seasons or sample sites.
Its use also facilitates interlaboratory comparisons.
11-5.8 Test temperature should be chosen based on conditions of particular interest, or to match the
conditions at the sample site. In either case, the choice of temperature and test organism should be compatible
(ASTM 1984). Suggested test temperatures may range from 7-33 ° C and should correspond to the average
spring-summer temperature of the study area (ASTM 1984).
11.5.9 Dissolved oxygen should be maintained between 40% and 100% saturation.
11.5.10 Light quality and daylength are important because of their impacts on both chemical degradation
and organism health. Light should be provided from cool-white fluorescent lamps at an intensity appropriate for
the test species (ASTM 1984).
11.5.11 The photoperiod can be selected to mimic that experienced at the sample site, or to simulate a
particular season. Suggested periods of daylight and darkness include 16 h light/8 h dark, 14 h light/10 h dark,
12 h light/12 h dark (13, ASTM 1984).
11.5.12 Whether or not test organisms should be fed during the test depends on test duration and the
type of test organism in use. The addition of food can complicate the interpretation of test results because it
adds new participate material, and the food may interact in unknown ways with contaminants in the sediments
(13). For acute tests (< 1 week) and many infaunal organisms which process sediments directly, enough
sediment has generally been provided to ensure adequate nutrition and feeding may not be necessary. If the
organisms are fish or filter feeders, food may be required, especially during long tests.
11.6 Chemical analysis of test water, sediment and organisms
11.6.1 Test water and sediments should be analyzed for contaminants of concern if the objectives of the
study are to determine the sources and concentrations of contaminants. If the test is designed to assess toxitity
only, then identification of sources of toxicity are not necessary.
11.6.2 Analyses of specific contaminants in tissues of the test organisms are needed if bioaccumulation
or bioconcentration is of interest. If measurement of organic chemicals, metals or other contaminants is
desireable, appropriate preservation methods should be followed when samples are collected.
12.0 Data interpretation
12.1 Bioaccumulation of contaminants or toxic effects such as mortality from sediment or sediment
extract exposure are important to the individuals of a particular species however, the ecological significance of
those data are difficult to predict (ref 12). Toxic effects observed in laboratory exposures may not reflect affects
on natural populations. However, bioaccumulation of a contaminant above a certain level or a toxicity response
higher (or lower) when compared to that same response in a population of organisms exposed to a control
sediment is undesirable.
122 The calculation procedure(s) and interpretation of the results should be appropriate to the
experimental design. Procedures used to calculate results of tests can be divided into two categories: those that
test hypotheses and those that provide point estimates. No procedure should be used without careful
consideration of (a) the advantages and disadvantages of various alternative procedures, and (b) appropriate
preliminary tests, such as those for outliers and for heterogeneity.
11
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123 When samples from field sites are independently replicated, site effects (bioaccumulation and
toxicity endpoints) can be statistically compared by t-tests, analysis of variance (ANOVA) or regression analysis.
Analysis of variance is used to determine whether any of the observed differences among the concentrations (or
samples) are statistically significant. This is a test of the null hypothesis that no differences exist in effects
observed among test concentrations (or samples) and controls. If the F-test is not statistically significant
(P>0.05), it can be concluded that the effects observed in the test material treatments (or field sites) were not
large enough to be detected as statistically significant by the experimental design and hypothesis test used. Non-
rejection does not mean that the null hypothesis is true. The NOEC based on this end point is then taken to be
the highest test concentration tested (33,34). The amount of effect that occurred at this concentration should be
considered.
123.1 All exposure concentration effects (or field sites) can be compared with the control effects by
using mean separation techniques such as those explained by Chew orthogonal contrasts (35), Fisher's methods,
Dunnett's procedure or Williams' method. The lowest concentration for which the difference in observed effect
exceeds the statistical significant difference is defined as the LOEC for that end point. The highest concentration
for which the difference in effect is not greater than the statistical significant difference is defined as the NOEC
for that end point (33).
12.4 In cases where sediment dilution series toxicity studies are conducted the LC50 or EC50 and its
95% confidence limits should be calculated (when appropriate) on the basis of (a) the measured initial
concentrations of test material, if available, or the calculated initial concentrations for static tests, and (b) the
average measured concentrations of test material, if available, or the calculated average concentrations for flow-
through tests. If other LC or ECs are calculated, their 95% confidence limits should also be calculated (see
ASTM E 729).
12.4.1 Most toxicity tests produce quantal data, that is, counts of the number of responses in two
mutually exclusive categories, such as alive or dead. A variety of methods (32) can be used to calculate an LC50
or EC50 and 95% confidence limits from a set of quantal data that is binomially distributed and contains two or
more concentrations at which the percent dead or effected is between zero and 100. The most widely used are
the probit, moving average, Spearman-Karber and Litchfield-Wilcoxon methods. The method used should
appropriately take into account the number of test organisms per chamber. The binomial test can also be used
to obtain statistically sound information about the LC50 or EC50 even when less than two concentrations kill or
affect between zero and 100 percent. The binomial test provides a range within which the LC50 or EC50 should
lie.
12
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop
ASTM Designing Sediment Tests
Table 1. Marine sediment toxicity tests.
TAXA
EXPOSURE REFERENCE
Mortality
Larval fish
Amphipods, bivalves
polychaetes, cumaceans
Amphipods
Fish bivalves
Fish
Shrimp, polychaetes
Shrimp So, Su
Copepods, amphipods, isopods,
shrimp
Amphipods, mysids, bivalves
fish
Mysids, amphipods
Phytoplankton
Fish
Shrimp, mussel, crab,
tunicate, lobster
El*
So"
So
Suc
So, Su
So
827,28
So, Su
So, Su
Su
El
El
Su
S7
818
819,21,22^9,60
and more
823
824
525,26
C17
Rogerson et al.
Gentile et al.
85,87,S&R
C15
C22.S7
S31
Avoidance /behavior
Echinoderm, lobster, crab,
shrimp, bivalve, amphipod
Amphipod
Crab, shrimp, fish, bivalve,
polychaete
Fish
Bivalves So
Polychaetes
Growth/reproduction/life cvcle
Fish
Bivalve Su
Mysids Su
Amphipods
Nematodes
Polychaetes
Copepods
Sea urchin
Polychaetes
Pathology
Fish
Bivalves, polychaetes, amphipods
Oyster, fish
So
So
So
So
813,14,16
So
S9
S10
Sll
S12
S15, Olla 1989
S4
Su
85
S17, G&al 85,87
Su S&R
So 832
So,Su S4,61
So C34
El C32
So, Su Johns et al
So
So, Su
Su,So
858, C30
Yevich et aL
Gardner et al.
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Table 1. Marine sediment toxicity test, (continued)
Physiology
Oligochaetes El CIO
Shrimp, polychaetes Su C35
Fish Su C36
Polychaetes So, Su Johns etal
Chromoome
Fish El S3,57,C9
Polychaetes Su Pesch et al.
Bacterial activity
BacteriaEl C12
BacteriaEl C13
Community
Macrobenthos So 83435,36
Macrobenthos So S37
Macrobenthos So S38
Macrobenthos So S33
Macrobenthos So S39
'El - elutriate, extract, pore water exposure
bSo - solid-phase sediment exposure
°Su - suspended sediment exposure
14
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop
ASTM Designing Sediment Tests
Table 2. Freshwater sediment toxicity tests.
TAXA
Mortality
Cladocerans, insect larvae,
isopods, fish, Urceus?
Insect larvae, dadocerans,
amphipods, fish
Cladocerans, insect larvae
Cladocerans
Cladocera, amphipods, insect larvae
Oligochaetes
Amphipods, insect larvae
Cladocerans
Insect larvae
Growth/reproduction
Insect larvae
Fish, dadocerans, bacteria,
Paratanytarsus
Insect larve, amphipods, dadocerans
Insect larvae
Nematodes
Physiology
Oligochaetes
Genetic damage
Fish
Nematodes
EXPOSURE
So
So, El
So
So, Su
So
So
So
So
So
So
So, El
So
So
El
El
El
El
REFERENCE
L1A3.4
L5
L6,15,16,17
L7
L8,9
L10,ll,12
L13
L14.20
L18
L18
L19
L9
L21,22
L29
L23,24
Ui5,26,27,28
L29
Bacterial activity
Bacteria El
Giesy et al.
15
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop MicrotoxR Elutriate Test
PERFORMING TOXICITY TESTS WITH THE MICROTOXR MODEL 500
I. GENERAL:
This document describes procedures for performing toxicity tests with the MicrotoxR Model
500. The instrument measures the light output of luminescent bacteria (supplied by the
manufacturer) before and after they are challenged by dilutions of a sample of unknown toxicity. A
Reagent Blank containing no toxicant is used to normalize the responses of the four sample test
concentrations during data reduction. The degree of light loss resulting from metabolic inhibition in
the test organisms indicates the toxicity of the sample and is used to determine a dose-response
curve from which the effective concentration of the sample is found. Measured light readings are
transmitted via a RS232 interface to a personal computer which estimates an Effective Concentration
using a data reduction program written in BASICR.
NOTE: Because individual users will select different computers and MicrobicsR periodically updates
its data reduction software, this document cannot provide detailed instructions for interfacing the
instrument with the computer or reducing data with the program supplied with the instrument.
Consult latest software guide for specific information.
II. REQUIRED REAGENTS
A. Microtox" Reconstitution Solution.
1. Distilled water, (may be stored indefinitely at room temperature).
B. Microtox" Osmotic Adjustment Solution (MOAS).
1. MOAS is a solution of de-ionized water containing 22% NaCl (220 ppt) used during
the standard bioassay procedure to osmotically adjust the sample, thereby
preventing cells from lysing. Generally, one part of MOAS is added to 10 parts of
sample. The assay is normally run at 2% NaCl. MOAS may be stored indefinitely
at room temperature.
C. Microtox" Diluent.
1. Diluent is 2% NaCl (20 ppt) used for diluting the sample and reagent. Diluent may
be stored indefinitely at room temperature.
D. Microtox" Reagent.
1. Reagent is a freeze-dried culture of a specially developed strain of the marine
bacterium Photobacterium phoshoreum. Reagent has a shelf-life of one year when
stored in a freezer at -20°C. SELF-DEFROSTING FREEZERS SHOULD NOT
BE USED FOR LONG TERM STORAGE OF THE REAGENT. Self defrosting
freezers periodically warm to prevent frost accumulation. Periodic warming of the
reagent may decrease storage tune and viability of the cultures.
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in. REQUIRED EQUIPMENT.
A. Microtox" disposable cuvettes
B. Pipettors and pipettor tips
1. 1, 10 /iL (white tips)
2. 1, 250 /iL (blue tips)
3. 1, 500 /iL (blue tips)
4. 1, Oxford 1000 /iL P-7000 Micropipettor (optional).
5. 1, Oxford/Nichiryo Model 8100 syringe dispenser and 15.0 mL syringes.
C. Microbics Microtox" Model 500
D. Microbics data capture and reduction program
E. Micro-computer with one serial port capable of running Micro Soft Basic or BasicA
software.
IV. INSTRUMENT PREPARATION.
A. To preform a single standard bioassay, place clean, unused cuvettes in Reagent Well and in
the incubator block wells in rows A and B.
B. Pipette 1 mL Reconstitution Solution into the cuvette in the Reagent Well
C. Pipette 500 /iL Diluent into each cuvette in wells Bl through B5
D. Pipette 1 mL Diluent into each cuvette in wells Al through A4
NOTE: To perform more than one assay at a time repeat steps A-D with additional cuvettes placed
hi rows C and D and E and F.
V. SAMPLE PREPARATION.
NOTE: A PRIMARY DILUTION OF THE SAMPLE MAY BE NECESSARY. REMEMBER
TO ACCOUNT FOR THE ADDITIONAL DILUTION IN DATA REDUCTION. (FOR DATA
OF OPTIMUM VALUE, TRY TO BRACKET THE ECSO WITH THE DILUTIONS)
A. Pipette 250 /iL MOAS into the cuvette in well A5.
B. Add 2.5 mL of sample (or Phenol Standard 90 mg/L) to cuvette A5, then mix by aspirating
and ejecting the sample, using the 500 /iL micropipettor.
C. Transfer 1.0 mL from A5 to A4, and mix two to three times as described above A4 using
the 500 /iL micropipettor.
D. Transfer 1.0 mL from A4 to A3, and mix A3 as described above using the 500 /iL
micropipettor.
E. Transfer 1.0 mL from A3 to A2, and mix A2 as described above using the 500 /iL
micropipettor.
F. Wait 5 minutes for solutions to come to controlled temperature.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Microtox" Elutriate Test
VI. REAGENT PREPARATION.
A. Just prior to reagent reconstitution, remove vial of MicrotoxR Reagent from the freezer.
Remove the seal and the stopper.
1. If the reagent pellet is not seated on the bottom of the vial, tap and shake the vial
until the pellet is seated.
B. Take the cuvette of reconstitution solution from the Reagent Well. Place the lip of the
cuvette on top of the reagent vial Then, as QUICKLY as possible, DUMP the
reconstitution solution into the reagent vial. Swirl the reagent into the reagent cuvette, put
the cuvette back in the reagent well.
C. Mix the reconstituted reagent 20 times by aspirating and ejecting the solution with a new tip
on the 500 /*L micropipettor.
D. Pipette 10 pL reconstituted reagent into the cuvette in wells Bl through B5.
NOTE: When transferring the 10 nL of reagent into a cuvette, leave both cuvettes in the wells.
Place the pipette tip under the surface of the liquid, but DO NOT REST THE PIPETTE TIP ON
THE BOTTOM OF THE CUVETTE.
SUGGESTION: Rest the 10 fiL pipette tip against the cuvette's inside rim. Slide the tip of
the pipet down until the ridge on the pipette tip touches the rim of the
cuvette. Stop there. The tip is in a good position for removing liquid from
the cuvette.
E. Mix the reagent in row B by aspirating and ejecting two to three times using a 250 pL
micropipettor.
F. Wait 15 minutes for reagent to stabilize.
NOTE: The reconstituted reagent is viable for approximately 2 hours.
VH. STANDARD ASSAY PROCEDURE.
A. Take the cuvette from well Bl, and place it in the turret well.
B. Press the SET button.
1. Wait for the Ready Green Light to illuminate on the front panel of the unit. DO
NOT PRESS THE SET BUTTON AGAIN FOR THIS ASSAY.
C. Read the initial (I0) light levels of the prepared cuvettes.
1. Place the cuvettes in the turret in the following order: Bl, B2, B3, B4, B5 and press
the read button after each.
a. The light reading for each cuvette will be displayed on the computer screen
in the appropriate sample number column.
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D. Make the following 500/iL transfers, mixing each sample by aspirating and ejecting 2-3 times
after each transfer using the 500 pL micropipettor: Al to Bl, A2 to B2, A3 to B3, A4 to B4,
A5 to B5.
1. The Gnal assay concentrations are approximately: 5.6, 11.3, 22.5 and 45 percent of
the sample being used. For example if the sample was originally a 100% sample
(undiluted) the resulting tested concentrations would be 5.5, 113, 22, and 45%.
However if the sample being tested was originally diluted 1:1 before the dilutions
were made to perform the assay, the tested concentrations would be 2.25, 5.65, 11,
and 22.6%.
E. When the final transfer and mixing is complete, HIT THE RETURN KEY.
1. Hitting the return key tells the computer to record how long the transfers took to
accomplish and display the elapsing time.
2. When TIME1 is elapsed the program will prompt you to transfer each curvet
(starting with Bl through B5) to the turret and push the READ button. The
prompt is the word "enter" which is displayed under each successive concentration.
The program spaces the prompts to accommodate for the time it took to make the
initial volume transfers between the A cuvettes and the B cuvettes.
3. When TIME2 has elapsed the program will again prompt you to transfer the
cuvettes starting with Al through B5 to the turret and press the READ button.
F. After the last reading is taken, the program will store data in the previously named data file
with either the TTME1 or TTME2 "dot designator" to differentiate between the two sets of
light readings.
Vffl. REDUCING STORED DATA WITH Microtox" DATA REDUCTION SOFTWARE.
A. Refer to the appropriate version of the MicrotoxR guide for detailed instruction on using the
data collection and reduction program.
DC. MICROTOX 100% ASSAY PROCEDURE.
NOTE: The standard Microtox" test procedure tests four sample dilutions, of which the highest
possible concentration is about 49.5%. Some Microtox users have a need to test samples
without dilution. At the present time, the automated data capture program cannot be used
with the 100% assay. Because the data capture program does not store the light readings it
is necessary to manually record the individual I0 and I, readings. The recorded data can be
used to generate an EC value using the data reduction program using the "Enter data from
the keyboard" option at the main menu.
This procedure differs from the standard assay in that initial light (I0) readings are
not recorded for each sample. To calculate an EC value using the data reduction
program, it is necessary to input the I0 reading obtained for the sample Al as the
"theoretical" I0 reading for each sample (Al through AS).
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Microtox" Elutriate Test
A. Instrument and sample preparation.
1. Add 1.0 mL of Reconstitution Solution to a cuvette in reagent well.
2. Add 1.0 mL diluent to cuvettes Al, A2, A3, and A4.
3. Add 50 mg AR grade Nad to A5.
4. Add 2.5 mL sample to A5, and mix A5 until the NaCl is dissolved. Discard 500 jiL
from A5.
5. Transfer 1.0 mL from A5 to A4 and mix A4.
6. Transfer 1.0 mL from A4 to A3, and mix A3.
7. Transfer 1.0 mL from A3 to A2, and mix A2.
8. Discard 1 mL from A2.
9. Set timer to 5 minutes to allow solutions to temperature equilibrate.
B. Reagent preparation and assay procedure.
1. Reconstitute a vial of Microtox" Reagent and mix 20 times with the 500 fiL
pipettor.
2. Start tuner set for 5 minutes (after the initial 5 minute temperature equilibration
period has passed). Transfer 10 /iL of reconstituted reagent to Al, A2, A3, A4, and
A5.
a. If using the 10 /iL pipettor to transfer reagent to the cuvettes, use pre-
cooled tips and discard each tip between transfers to prevent contaminating
Reagent stock.
b. If using the Oxford/Nichiryo model 8100 multiple pipettor, use a syringe
which has been pre-cooled by placing in a refrigerator or held in a beaker
containing ice (pre-cooling the syringe minimizes wanning of reagent while
it is in the dispenser).
3. Mix each cuvette from Al through A5 using a 500 pL pipettor. Record the time
required to complete the transfers of the Reagent to the cuvettes and divide the
time in seconds by 5. The resulting time will be the interval between each reading
at the five and fifteen minute readings. Normally, the time between transfers is
accounted for by the automated program.
4. At five minutes after beginning of reagent transfer, place the Al cuvette in the
turret and press the SET button.
5. After the instrument is set, press the read button with Al in the turret and record
the value.
6. Read the remaining samples at the appropriate interval by placing each in the turret
and pressing the READ button. Record each value for subsequent data reduction.
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7. Beginning at 15 minutes, re-read the samples starting with Al proceeding through
A5 allowing for the appropriate tune interval between readings as calculated in step
2 (above). Record all values for data reduction..
C. Data reduction of 100% assay.
1. Refer to the appropriate version of the MicrotoxR guide for detailed instruction on
using the data collection and reduction program as it applies to the 100% assay.
X. COLOR CORRECTION PROCEDURE.
NOTE: Colored aqueous samples, particularly those colored red or brown may cause non-specific
reductions in light level when analyzed according to the standard Microtox assay procedure.
These light level reductions cannot be distinguished from those caused by toxicants in the
standard toxicity assay. The following procedure, utilizing a special Color Correction
Cuvette, measures the amount of color interference in a given sample. The measurement is
then used to correct the results obtained for the sample. Reconstituted Reagent left over
from the standard toxicity assay may be used for this procedure even if it is several hours
old.
The color correction procedure is necessary only when appreciable color is visible in
a diluted sample near the EC50 concentration.
A. Sample Preparation.
1. If the sample is turbid, centrifuge at 10,000 x G for 15 minutes.
2. Perform standard assay, and determine the EC50. If the EC50 concentration has no
visible color, the color correction is not required.
3. If color is appreciable at the EC50 concentration, make a sample dilution (mm. vol.
2 mL.) dose to the ECSO concentration, (example EC50 = 7.5%, make a 5% or
10% sample)
4. Add 2.0 mL of the diluted sample to cuvette in A4.
5. If more than one sample is to be run at this time, repeat steps 3 and 4 above for
each sample using B and C wells for storage. The sequence of sample reading
should be "least color" first to "most color" last.
B. Instrument Preparation.
1. Add 1.0 mL Diluent to cuvette in A2.
2. Add 2.0 mL Diluent to cuvette in A5 and OUTER CHAMBER of the Color
Correction Cuvette in Al.
3. Wait 5 minutes.
C. Color Correction Procedure.
1. Transfer 50 /iL reagent to A2 and mix using 500 /iL pipettor.
2. Using a glass Pasteur pipette, transfer diluted reagent to center chamber of the
Color Correction Cuvette until the reagent is at diluent level.
3. Place Color Correction Cuvette in turret. Wait 5 minutes. Press SET button.
4. Wait for ready green light then press the READ button. Record the first blank
light level reading (B0).
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Microtox" Elutriate Test
CAUTION: Do not move or rotate the Color Correction Cuvette until the entire color
correction procedure has been completed.
5. Immediately remove and discard diluent from outside chamber of the Color
Correction Cuvette, using a glass Pasteur pipette.
6. Transfer entire volume of prepared diluted sample (from A4) to outside chamber of
Color Correction Cuvette using the Pasteur pipette.
7. Five minutes after the B0 reading (Time 0) press READ and record the light level
IT.
8. Remove and discard sample from outside chamber of Color Correction Cuvette
using an aspirator. If you have additional colored samples, read their light levels at
this tune, repeating steps 6 and 7 (above) for each sample. Time the readings at 5
minute intervals, T0 + 10, T0 + 15, T0 + 20, etc...
9. Transfer entire volume of diluent in A5 to outside chamber of Color Correction
Cuvette using a Pasteur pipette.
10. Wait 5 minutes from sample reading, press READ button. Record second blank
light level reading.
D. Tabulating and reducing color correction data.
1. Consult the appropriate version of the Microtox" guide for detailed instruction on
using the data collection and reduction program to correct for sample color.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Daphnia magna Elutriate Testing
Daphnia magna Elutriate Testing Experimental Design
Static acute toxicity tests will be conducted with Daphnia magna and sediment elutriates. The
daphnids will be exposed for 48 hours to full strength (100%) elutriates and 50%, 25%, and 12.5% dilutions
of the 100% elutriates, and to a dilution water control without replication. Dilutions will be prepared a fresh
water.
Ten Daphnia magna (<24 hours old) will be placed into each 250 mL test beaker in 200 mL of test
solutions. Adult daphnids will be isolated from laboratory cultures on Day -1 of the test. Young daphnids
(<24-hours old) will be removed from the cultures on Day 0 and placed into a culture water box. Ten of the
young daphnids will be removed from the culture water box with smooth glass tubes (large bore) and placed
directly into each 250-mL test beaker in the order of dilution water controls, 12^%, 25%, 50%, and 100%
sediment elutriates. The test temperature will be maintained at 20°C with a temperature-controlled
waterbath. The photoperiod for the tests will be 16:8 (light:darkness) with a light intensity of about 50 fc.
The daphnids will not be fed during the tests.
The pH, total hardness, alkalinity, conductivity, ammonia, dissolved oxygen, turbidity, chloride, and
sulfate will be determined on the fresh water, and on each 100% elutriate (except sulfate) sample. On Day 0
of the test, pH, dissolved oxygen, and conductivity will be measured in the 100% elutriate samples before
dilutions are made. Dissolved oxygen in the 100% elutriate samples will be adjusted at this time if necessary.
At the end of each test, pH, dissolved oxygen, and conductivity will be measured in the 100%, 25%, and 0%
elutriate treatments.
Survival of the daphnids will be recorded in all treatments at 24 and 48 hours. The lack of mobility
in response to prodding with a blunt probe during 5 seconds of observation will be used as criteria to
determine death.
I. Type of test.
A. 48-hour acute toxicity test.
B. Toxicants.
1. Sediment elutriates.
C. Test conditions.
1. Fresh water dilutions of the sediment elutriates.
2. Temperature: 20°C
3. No Feed.
4. Dilution water quality.
a. Fresh water 134 mg/L total hardness as CaCO3, alkalinity 65 mg/L CaCO3, sulfate 72
mg/L, pH 73, conductivity 245 /mhos.
5. Photoperiod of 16:8 (light:dark) with light intensity of about 50 fc.
6. Ten daphnids per test chamber, no replication.
n. Test description.
A. Treatments.
1. Freshwater control.
2. 100% sediment elutriate.
3. 50% sediment elutriate.
4. 25% sediment elutriate.
5. 115% sediment elutriate.
B. Treatments will not be replicated.
C. 200-mL test solution per 250-mL test beaker.
D. 375-mL sediment elutriate per test.
III. Pre-test Preparation.
A. Regulate water bath temperature to 20°C.
B. Run Microtox on sediment elutriates if storage tune exceeds two weeks.
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IV. Water quality monitoring.
A. pH, total hardness, alkalinity, conductivity, ammonia, dissolved oxygen, turbidity, chloride, and sulfate
on fresh water.
B. pH, alkalinity, total hardness, conductivity, oxygen, chloride, ammonia, and turbidity on each elutriate
sample.
C. pH, dissolved oxygen, and conductivity on the elutriate samples on Day 0 of the test before dilutions
are made.
D. pH, dissolved oxygen, and conductivity in the 100%, 25%, and 0% elutriate dilutions at the end of
the test.
E. Temperature will be monitored daily in the test waterbaths.
V. Test stocking regime.
A. Ten .<. 24-hour old Daphnia mflg"? per test beaker.
B. Count groups of 5 daphnids directly into 250-mL test beakers until there is a total of 10 per beaker.
VI. Biological Sampling.
A. Survival will be recorded in all treatments at 24 and 48 hours.
B. Lack of mobility hi response to prodding with a blunt probe during 5 seconds of observation will be
used as criteria to determine death.
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Nelson, Coylc and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
STANDARD GUIDE FOR CONDUCTING SEDIMENT TOXICITY TESTS
WITH FRESHWATER INVERTEBRATES1
M.K. Nelson, C.G. Ingersoll, and FJ. Dwyer
U.S. Fish and Wildlife Service
National Fisheries Contaminant Research Center
Route 2, 4200 New Haven Road
Columbia, MO 65201
(314)-875-5399 FTS 276-1800
1. Scope
1.1 This guide describes procedures for obtaining laboratory data to evaluate adverse effects of contaminants
associated with whole sediment on freshwater organisms. The methods are designed to assess the toxic effects on
invertebrate survival, growth, or reproduction, from short (for example, 10 days) or long-term tests, in static or
flow-through water systems. Sediments to be tested may be collected from field sites or spiked with known
compounds in the laboratory. Test procedures are described for three species, (1) Hvalella azteca. (2)
Cbironomus tentans. and (3) Chironomus riparius. Methods described in this document should also be useful for
conducting sediment toxicity tests with other aquatic species, although modifications may be necessary.
12 Modification of these procedures might be justified by special needs. Results of tests conducted using
unusual procedures are not likely to be comparable to results using this guide. Comparison of results obtained
using modified and unmodified versions of these procedures might provide useful information concerning new
concepts and procedures for conducting sediment toxiciry tests with freshwater organisms.
13 The results from field collected sediments used in toxitity tests to determine a spatial or temporal
distribution of sediment toxicity may be reported in terms of the biological effects on survival, growth, or
reproduction (see Section 16, Calculation of Results). In addition, these procedures are applicable to most
sediments or chemicals added to sediment. Materials either adhering to sediment particles or dissolved in
interstitial water can be tested. With appropriate modifications these procedures can be used to conduct
sediment toxicity tests when factors such as temperature, dissolved oxygen, pH, and sediment characteristics (for
example, particle size, organic carbon content, total solids) are of interest, or when there is a need to test such
materials such as sewage sludge, oils and paniculate matter. These methods might also be useful for conducting
bioaccumulation tests.
1.4 Results of toxicity tests with test materials experimentally added to sediments may be reported in terms
of an LC50 (median lethal concentration), and sometimes an EC50 (median effect concentration). Results of
tests may be reported in terms of an NOEC (no observed effect concentration) and LOEC (lowest observed
effect concentration).
1 This test method is under the jurisdiction of ASTM Committee E-47 on Biological Effects and
Environmental Fate and is the direct responsibility of Subcommittee E-47.03 on Sediment Toxicity.
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1_5 This guide is arranged as follows:
Referenced Documents 2
Terminology 3
Summary of Guide 4
Significance and Use 5
Interferences 6
Safety Precautions 7
Apparatus 8
Overlying Water 9
Sediment Characterization 10
Test Organisms 11
Experimental Design 12
Procedure 13
Analytical Methodology 14
Acceptability of Test 15
Calculation of Results 16
Documentation 17
Annexes
XI. Hvalella azteca (Amphipoda)
X2. <^hironom\}§ t?"t8IHS (Diptera)
X3. Chironomus riparius (Diptera)
1.6 This guide addresses procedures which may involve hazardous materials, operations, and equipment, and
it does not purport to address all of the safety problems associated with iu use. It is the responsibility of the user
to establish appropriate safety and health pracdces, and determine the applicabih'ty of regulatory limitations prior
to use. While some safety considerations are included in this document, it is beyond the scope of this document
to encompass all safety requirements necessary to conduct sediment toxitity tests. Precautionary statements are
given in Section 7.
2. Applicable Documents
2.1 ASTM Standards:
E 380 Standard for Metric Practice2
E 729 Practice for Conducting Acute Toxicity Tests with Fishes, Macroinvertebrates, and Amphibians3
E 943 Standard Definitions of Terms Relating to Biological Effects and Environmental FateS
E 1023 Guide for Assessing the Hazard of a Material to Aquatic Organisms and Their Uses3
E 1241 Guide for Conducting Early Life-Stage Toxiciry Tests with Fishes3
D 1129 Definitions of Terms Relating to Water4
D 4387 Guide for Selecting Grab Sampling Devices for Collecting Benthic Macroinveitebrates3
D 4447 Guide for Disposal of Laboratory Chemicals and Samples3
2 Annual Book of ASTM Standards, Vol 14.02.
3 Annual Book of ASTM Standards, Vol 11.04.
4 Annual Book of ASTM Standards, Vol 11.01.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
D 4823 Guide for Core Sampling Submerged, Unconsolidated Sediments5
E 1367 Standard Guide for Conducting Solid Phase 10-Day Static Sediment Toxicity Tests with Marine and
Estuarine Infaunal Amphipods
E 1391 Standard Guide for Collection, Storage, Characterization, and Manipulation of Sediments for
lexicological Testing
E XXX Guide for Designing Sediment Toxicity Tests (Draft 4)
3. Terminology
3.1 The words "must", "should", "may", "can", and "might" have very specific meanings in this guide. "Must" is
used to express an absolute requirement, that is, to state that the test ought to be designed to satisfy the specified
condition, unless the purpose of the test requires a different design. "Must" is only used in connection with the
factors that directly relate to the acceptability of the test (see Section 15). "Should" is used to state that the
specified condition is recommended and ought to be met if possible. Although a violation of one "should" is
rarely a serious matter, violation of several will often render the results questionable. Terms such as "is
desirable", "is often desirable", and "might be desirable" are used in connection with less important factors. "May
is used to mean "is(are) allowed to", "can" is used to mean "is(are) able to", and "might" is used to mean "could
possibly". Thus, the classic distinction between "may" and "can" is preserved, and "might" is never used as a
synonym for either "may" or "can".
32 Descriptions of Terms Specific to this Guide:
32.1 sediment—a naturally occurring paniculate material which has been transported and deposited at the
bottom of a body of water, or an experimentally prepared substrate within which the test organisms can interact.
3.22 whole sediment—distinguished from elutriates, and resuspended sediments, in that the whole, intact
sediment is used to expose the organisms, not a form or derivative of the sediment.
3.23 clean—denotes a sediment or water that does not contain concentrations of test materials which cause
apparent stress to the test organisms or reduce their survival
32.4 overlying water—the water placed over the whole sediment in the test chamber for the conduct of the
toxicity test, and may also include the water used to manipulate the sediments.
32.5 interstitial water—the water within a wet sediment that surrounds the sediment particles, expressed as
the percent ratio of the weight of the water in the sediment to the weight of the wet sediment.
32.6 spiking—the experimental addition of a test material such as a chemical or mixture of chemicals,
sewage sludge, oil, paniculate matter, or highly contaminated sediment to a dean negative control or reference
sediment, such that the toxicity of the material added can be determined. After the test material is added, which
may involve a solvent carrier, the sediment is thoroughly mixed to evenly distribute the test material throughout
the sediment.
32.7 concentration—the ratio of weight or volume of test material(s) to the weight or volume of sediment.
5 Annual Book of ASTM Standards, Vol 11.02.
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33 For definitions of other terms used in this guide, refer to Standards E 729, E 943, E 1023, E 1241 and D
1129. For an explanation of units and symbols, refer to Standard £ 380.
4. Summary of Guide
4.1 The toxicity of contaminated whole sediments is assessed during continuous exposure of aquatic
organisms, using either static or flow-through exposure systems. Sediments tested may either be collected from
field sites or spiked with a known compound(s). A negative control sediment or a reference sediment is used to
(a) give a measure of the acceptability of the test, (b) provide evidence of the health and relative quality of the
test organisms, (c) determine the suitability of the overlying water, test conditions, food, handling procedures, and
(d) provide a basis for interpreting data obtained from the test sediments. A reference sediment is collected
from the field in a dean area and represents the test sediments in sediment characteristics (for example, TOC,
parades size, pH). Specified data are obtained to determine the toxic effects on survival, growth, or
reproduction, from short (for example, 10 days), or long-term exposures to aquatic invertebrates.
5. Significance and Use
5.1 Protection of a spedes requires averting detrimental contaminant related effects on the survival, growth,
reproduction, health, and uses of the individuals of that spedes (1). Sediment toxicity tests provide information
concerning the bioavailability of contaminants associated with sediments to aquatic organisms. Invertebrates
occupy an essential niche in aquatic ecosystems and are an important food source for fish, wildlife, and larger
invertebrates. A major change hi the availability of invertebrates as either a food source, or as organisms
functioning properly in trophic energy transfer and nutrient cycling, could have serious adverse ecological effects
on the entire aquatic system.
5.2 Results from sediment toxicity tests might be an important consideration when assessing the hazards of
materials on aquatic organisms (see Guide E 1023) or when deriving sediment quality concentrations for aquatic
organisms (2).
53 Information might also be obtained on accumulation of contaminants assodated with sediments by
analysis of animal tissues for the contaminant(s) being monitored.
5.4 The sediment toxicity test might be used to determine the temporal or spatial distribution of sediment
toxicity. Test methods can be used to detect horizontal and vertical gradients in toxicity.
55 Results of sediment toxicity tests with test materials experimentally added to sediments could be used to
compare the sensitivities of different spedes, the toxicity of different test materials, and to study the effects of
various environmental factors or results of such tests. Results of sediment toxicity tests are useful for studying
biological availability of test materials, and structure-activity relationships.
5.6 Results of sediment toxicity tests can be used to predict effects likely to occur with aquatic organisms in
field situations as a result of exposure under comparable conditions, except that (a) motile organisms might avoid
exposure and (b) toxidty to benthic organisms can be dependent on sediment physical characteristics, dynamics of
equilibrium partitioning, and the route of exposure.
5.6.1 Field surveys can be designed to provide either a qualitative reconnaissance of the distribution of
sediment toxicity or a quantitative statistical comparison of toxidty among sites.
5.6.2 Sediment toxidty surveys are usually part of more comprehensive analyses of biological, chemical,
geological, and hydrographic conditions. Statistical correlation can be improved and costs reduced if subsamples
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
for sediment toxicity tests, geochemical analyses, and benthic community structure are taken simultaneously from
the same grab of the same site.
5.7 Sediment toxicity tests can be an important tool for making decisions regarding the extent of remedial
action needed for contaminated aquatic sites.
6. Interferences
6.1 Limitations to the methods described in this guide might arise and thereby influence sediment toxicity
test results and complicate data interpretation. The following factors should be considered when testing whole
sediments:
6.1.1 Alteration of field samples in preparation for laboratory testing (for example, sieving).
6.1.1.1 Maintaining the integrity of the sediment environment during its removal, transport, and testing in the
laboratory is extremely difficult. The sediment environment is composed of a myriad of microenvironments,
redox gradients and other interacting physiochemical and biological processes. Many of these characteristics
influence sediment toxicity and bioavailability to benthic and planktonic organisms, microbial degradation, and
chemical sorption. Any disruption of this environment complicates interpretations of treatment effects, causative
factors, and jn situ comparisons.
6.1.1.2 Sediments tested at temperatures other than what they are collected might affect contaminant
solubility, partitioning coefficients, and other physical and chemical characteristics.
6.1.2 Interaction between sediment and overlying water and the influences of the ratio of sediment to
overlying water.
6.1.3 Interaction among chemicals present in the sediment.
6.1.4 Use of laboratory spiked sediment that might not be representative of contaminants associated with
sediments in the field.
6.1.5 Maintenance of acceptable quality of overlying water.
6.1.6 Addition of food (3) or solvents to the test chambers might obscure the adverse influence of
contaminants associated with sediment, provide an organic substrate for bacterial or fungal growth, and might
affect water quality characteristics^).
6.1.7 Resuspension of sediment during the toxicity test
6.1.8 Natural geochemical properties of test sediment collected from the field might not be within the
tolerance limits of the test species,
6.1.9 Recovery of test organisms from the sediment,
6.1.10 Field collected sediments may contain indigenous organisms including (a) predators, (b) the same or
closely related species to that being tested, and (c) microorganisms (for example, bacteria and molds) and algae
species that might grow in or on the sediment and test chamber surfaces.
6.L11 Test material concentrations might be reduced in the overlying water in flow-through testing, and
compounds such as ammonia might increase during testing.
62 Static tests might not be applicable with materials that are highly volatile or rapidly transform
biologically or chemically. The dynamics of test material partitioning between solid and dissolved phases at the
start of the test should therefore be considered, especially in relation to assumptions of chemical equilibria.
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7. Safety Precautions
7.1 Many substances pose health risks to humans if adequate precautions are not taken. Information on
toxitity to humans, recommended handling procedures, and chemical and physical properties of the test material
should be studied before a test is begun and made aware to all personnel involved (5,6,7,8). Contact with st
materials, overlying water and sediments should be minimized.
7.1.1 Many materials can adversely affect humans if precautions are inadequate. Skin contact with test
materials and solutions should be minimized by such means as wearing appropriate protective gloves, labt . ..iory
coats, aprons, and safety glasses, and by using dip nets, sieves or tubes to remove test organises from overlying
water. When handling hazardous sediments the proper handling procedures might include (a) sieving and
distributing sediments under a ventilated hood or in an enclosed glove box, (b) enclosing and ventilating the
tenacity testing water bath, and (c) use of respirators, aprons, safety glasses, and gloves. Field collected s; ments
might contain tor c materials and should be treated wii1- aution to minimize occupational exposure to workers.
Worker safety should also be considered when working with spiked sediments containing organics or inorganic
r:-<3'-<- those that are radio-labeled, and ith materials i are, or -e suspected of being, cartinoecnic
- Careful consideration should be given '• : hose cnem»< is which might biodegrade, trar. o more
toxic components, volatilize, oxidize, or photolyze during the test period.
13 For tests involving spiked sediments with known test materials, removal or degradation of test material
before disposal of stock solutions, overlying water, and sediments is sometimes desirable.
" alth and safetv precautions and applicable regulations for disposal of stock solutvis, test organisms,
sedimenu, and overlying water should be considered before beginning a test (ASTM D 4447).
15 Cleaning of equipment with a volatile solvent such as acetone should be performed only in a well-
ventilated area in which no smoking is allo'- - ' and no open flame such as a pilot light is present.
7.6 An acidic solution 'h not be mixtd with a hypochlorite solution because hazardous fumes might be
producec.
7.7 To prepare dilute acid solutions, concentrated acid should be added to water, not vice versa. Opening a
bottle of concer red acid and adding concentrated acid to water should be performed ooh in a fume hood.
7.8 Use of ground fault systems and leak detectors is strongly recommended to help prevent electrical
shocks.
8. Apparauis
8.1 Facilities - The facility should include constant temperature areas for culturing and testing to reduce
the possibility of contamination by test materials and other substances, especially volatile compounds. Holding,
acclimation, and culture tanks should not be in a room in which toxicity tests are conducted, stock solutions or
test solutions are prepared, or equipment is cleaned Test chambers may be placed hi a temperature controlled
recirculating water bath or a constant-temperature area. Air used for aeration should be free of fumes, on. and
water. Filters to remove oil, water, and bacteria are desirable. Air filtration through a 0.22 pm bacteria: .liter or
other suitable system may be used. The test facility should be well ventilated and free of fumes. Enclosures
might be desirable to ventilate icst chambers.
8.1.2 If a photoperiod other than continuous light is used, a timing device should be used to provide a
lightdaikness cycle. A 15- to 30-minute transition period (9) when lights go on and off it might be desirable to
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
reduce the possibility of test organisms being stressed by instantaneous illumination; a transition period when
lights go off might also be desirable.
8.2 Constructing Mfltgnals — Equipment and facilities that contact stock solutions, test solutions, sediment
and overlying water, into which test organisms will be placed, should not contain substances that can be leached
or dissolved in amounts that adversely affect the test organisms. In addition, equipment and facilities that contact
sediment or water should be chosen to minimi?? soiption of test materials from water. Glass, type 316 stainless
steel, nylon, high density polyethylene, polycarbonate and fluorocarbon plastics should be used whenever possible
to minimi7p. leaching, dissolution, and sorption. Concrete and rigid (unplasticized) plastics may be used for
holding, acclimation, and culture tanks, and in the water-supply system, but these materials should be soaked,
preferably hi flowing water, for a week or more before use (10). Cast-iron pipe should probably not be used in
freshwater-supply system because colloidal iron will be added to the overlying water and strainers will be needed
to remove rust particles. Copper, brass, lead, galvanized metal, and natural rubber should not contact overlying
water or stock solutions before or during the test. Items made of neoprene rubber and other materials not
mentioned above should not be used unless it has been shown that their use will not adversely affect survival,
growth, or reproduction of the test organisms.
83 Water Delivery System -- The water delivery system used hi flow-through testing can be one of several
designs. The system should be capable of delivering water to each replicate test chamber. Several designs of
diluter systems are currently in use; Mount and Brungs (11) diluters have been successfully modified for
sediment testing and other diluter systems have also been useful according to Ingersoll and Nelson (4) and Maki
(12). Various metering systems, using different combinations of siphons, pumps, solenoids, valves, etc., have been
used successfully to control the flow rates of overlying water.
83.1 The metering system should be calibrated before the test by determining the flow rate of the overlying
water through each test chamber. The general operation of the metering system should be visually checked daily
throughout the conduct of the test. If necessary the water delivery system should be adjusted during the test. At
any particular tune during the test, flow rates through any two test chambers should not differ by more than 10%.
8.4
8.4.1 In a toxicity test with aquatic organisms, test chambers are defined as the smallest physical units
between which there are no water connections. However, screens, cups, etc, may be used to create two or more
compartments within each chamber. Therefore, the overlying water can flow from one compartment to another
within a test chamber but, by definition, cannot flow from one chamber to another. All test chambers and
compartments if used, in a sediment toxicity test, must be identical. For the static tests, cover watch glasses may
be used to fit over the top of the test chambers such that an aeration dp is accommodated.
8.4.2 Test chambers may be constructed in several ways of various materials, depending on the experimental
design and the contaminants of interest Clear silicone adhesives, suitable for aquaria, sorb some organic
compounds which might be difficult to remove. Therefore, as little adhesive as possible should be hi contact with
test solution. If extra beads of adhesive are needed, they should be on the outside of the test chambers rather
than on the inside. To leach potentially toxic compounds from the adhesive, all new test chambers constructed
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using silicone adhesives should be acclimated at least 48 hours in overlying water used in the sediment toxitity
test.
8.43 Species-specific information on test chambers is given in each appendix (see Species Specific
Appendices).
8.5 f-lefli^ng ~ Test chambers, water delivery systems, equipment used to prepare and store overlying water,
and stock solutions, should be cleaned before use. New items should be washed in the following manner: (a)
detergent wash, (b) water rinse, (c) water-miscible organic solvent wash, (d) water rinse, (e) acid wash (such as
10% concentrated hydrochloric acid), and (f) rinsed at least twice with distilled, deionized, or overlying water.
Test chambers should be rinsed with overlying water just before use.
8.5.1 Many organic solvents leave a film that is insoluble in water. A dichromate-sulfuric acid cleaning
solution can generally be used in place of both the organic solvent and the acid (see ASTM E 729), but the
solution might attack silicone adhesive and leave chromium residues on glass.
8 52 Upon completion of a test, all items to be used again should be immediately (a) emptied of sediment
and overlying water (and properly disposed), (b) rinsed with water, (c) cleaned by a procedure appropriate for
removing the test material (for example, acid to remove metals and bases; detergent, organic solvent, or activated
carbon to remove organic chemicals), and (d) rinsed at least twice with distilled, deionized, or overlying water.
8.6 Acceptability ~ Before a toxicity test is conducted in new test facilities, it is desirable to conduct a "non-
toxicant" test, in which all test chambers contain a negative control or reference sediment, and overlying water
with no added test material Survival, growth, or reproduction of the test species will demonstrate whether
facilities, water, control sediment, and handling techniques are adequate to result in acceptable species-specific
control numbers. The magnitude of the within-chamber and between-chamber variance should also be
determined.
9. Overlying Water
9.1 Requirements - Besides being available in adequate supply, overlying water used in toxicity tests, and
water used to hold organisms before testing, should be acceptable to test species and uniform in quality. To be
acceptable to the test species, the water must allow satisfactory survival and growth, without showing signs of
disease or apparent stress, such as discoloration, or unusual behavior.
92 Source
9.2.1. Natural overlying water should be uncontaminated and of constant quality and should meet the
following specifications as established in ASTM E 729. The values stated help to ensure that test organisms are
not apparently stressed during holding, acclimation, and testing, and that test results are not unnecessarily
affected by water quality characteristics:
Paniculate matter <5 mg/L
TOC <5 mg/L
COD <5 mg/L
Residual chlorine < 11
9.2.1.2 A natural overlying water is considered to be of uniform quality if the monthly ranges of the
hardness, alkalinity, and specific conductance are less than 10% of their respective averages and if the monthly
range of pH is less than 0.4 unit Natural overlying waters should be obtained from an uncontaminated well or
spring, if possible, or from a surface water source. If surface water is used, the intake should be positioned to
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
minimize fluctuations in quality and the possibility of contamination and maximize the concentration of dissolved
oxygen and to help ensure low concentrations of sulfide and iron. Municipal water supplies often contain
unacceptably high concentrations of copper, lead, zinc, fluoride, chlorine or chloramines, and quality is often
variable (13). Chlorinated water should not be used for, or in the preparation of, overlying water because
residual chlorine and chlorine-produced oxidants are toxic to many aquatic animals (14). Dechlorinated water
should only be used as a last resort, because dechlorination is often incomplete.
9.2.2 For certain applications the experimental design might require use of water from the test sediment
collection site.
9.23 Reconstituted water is prepared by adding specified amounts of reagent grade8 chemicals to high
quality distilled or deionized water (see ASTM E 729). Acceptable water can be prepared using deionization,
distillation, or reverse-osmosis units. Conductivity, pH, hardness and alkalinity should be measured on each
batch of reconstituted water. If the water is prepared from a surface water, total organic carbon or chemical
oxygen demand should be measured on each batch. Filtration through sand, rock, bag, or depth-type cartridge
filters may be used to keep the concentration of paniculate matter acceptably low. The reconstituted water
should be intensively aerated before use, except that buffered soft fresh waters should be aerated before, but not
after, addition of buffers. Problems have been encountered with some species in some fresh reconstituted waters,
but these problems can be overcome by aging the reconstituted water for one or more weeks.
93 Characterization — The following items should be measured at least twice each year, and more often if
(a) such measurements have not been determined semiannually for at least two years, or (b) if surface water is
used:
93.1 pH, paniculate matter, TOC, organophosphorus pesticides, organic chlorine (or organochlorine
pesticides plus PCBs), chlorinated phenoxy herbicides, ammonia, cyanide, sulfide, bromide, chloride, fluoride,
iodide, nitrate, phosphate, sulfate, calcium, magnesium, sodium, potassium, aluminum, arsenic, beryllium, boron,
cadmium, chromium, cobalt, copper, iron, lead, manganese, mercury, molybdenum, nickel, selenium, silver, and
zinc, hardness, alkalinity, and conductivity (see ASTM E 729).
932 For each method used the detection limit should be below (a) the concentration in the overlying water,
or (b) the lowest concentration that has been shown to adversely affect the test species (14).
933 Water that might be contaminated with facultative pathogens may be passed through a properly
maintained ultraviolet sterilizer (15) equipped with an intensity meter and flow controls or passed through a filter
with a pore size of 0.45 ftm or less.
93.4 Water might need intense aeration using air stones, surface aerators, or column aerators (16,17,18).
Adequate aeration will stabilize pH, bring concentrations of dissolved oxygen and other gases into equilibrium
with air, and minimize oxygen demand and concentrations of volatiles. The concentration of dissolved oxygen in
6 "Reagent Chemicals, American Chemical Society Specifications," Am. Chemical Soc., Washington, DC.
For suggestions on the testing of reagents not listed by the American Chemical Society, see "Reagent
Chemicals and Standards," by Joseph Rosin, D. Van Nostrand Co., Inc., New York, NY, and the "United
States Pharmacopeia."
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water should be between 90% and 100% saturation (19) to help ensure that dissolved oxygen concentrations are
acceptable in test chambers.
10. Sediment Characterization
10.1 General — Before the preparation or collection of sediment an approved written procedure should be
prepared for the handling of sediments which might contain unknown quantities of toxic contaminants (see
Section 7, Safety Precautions). All sediments should be characterized and at least the following determined: pH,
organic carbon content (total organic carbon TOC) or total volatile sulfides, particle size distribution (percent
sand, silt, clay), and percent water content (20,21). Other analyses on sediments might include biological oxygen
demand, chemical oxygen demand, cation exchange capacity, Eh, pE, total inorganic carbon, total volatile solids,
acid volatile sulfides, total ammonia, metals, organosilicones, synthetic organic compounds, oil and grease,
petroleum hydrocarbons, and interstitial water analysis. Macrobenthos may be determined by a subsample of the
field collected sediment, lexicological results might provide information directing a more intensive analysis.
Sediment toxicity testing procedures are detailed in Section 13, Procedures.
10.2 Negative Control and Reference Sediment — A negative control sediment or a reference sediment is
used to (a) give a measure of the acceptability of the test, (b) provide evidence of the health and relative quality
of the test organisms, (c) determine the suitability of the overlying water, test conditions, food, handling
procedures, and (d) provide a basis for interpreting data obtained from the test sediments. Every test requires a
negative sediment control (sediment known to be non-toxic to, and within the geochemical requirements of the
test species) or a reference sediment. A reference sediment should be collected from the field in a dean area
and represent the test sediment in sediment characteristics (for example, TOC, particles size, pH). This provides
a site-specific basis for comparison of toxic and non-toxic conditions. The same overlying water, conditions,
procedures, and organisms should be used as in the other treatments, except that none of the test material(s)
being tested, or contaminated field collected sediments, is added to the negative control or reference sediment
test chambers.
10.2.1 If a field sediment has properties such as, grain size and organic content which might exceed the
tolerance range of the test species, it is desirable to include a reference sediment for these characteristics.
10J Field Collected Test Sediment
103.1 Collection (see Section 7, Safety Precautions). A benthic grab or core should be used rather than a
dredge to minimize disruption of the sample (see ASTM Guide for Collection, Storage, Characterization, and
Manipulation of Sediments for Toxicological Testing and ASTM Standard Guide D 4387). If the sediment is
obtained with a grab, it is preferable to collect a sediment sample from the upper 2 cm. This operation is
facilitated if the grab can be opened from the top so that the undisturbed sediment surface is exposed. The
sample should be transferred to a clean (see Section 8.5, Cleaning) sample container. If the contaminants
associated with sediments include compounds that readily photolyze, minimise direct sunlight during collection.
All sediment samples should be cooled to 4°C ±2°C in the field.
1032 Storage. Sediment samples should be stored at 4°C ±2°C and for no longer than two weeks before
the start of the test Freezing and longer storage might change sediment properties and should be avoided (see
ASTM Guide for Collection, Storage, Characterization, and Manipulation of Sediments for Toxicological
Testing). Sediment may be stored in containers constructed of suitable quality as outlined in Section 82,
Construction Materials. It is desirable to avoid contact with metals, including stainless steel and brass sieving
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screens, and some plastics. The samples should be thoroughly mixed and may be wet-press sieved through a
suitably sized sieve to remove large particles and indigenous organisms, especially predators. Sediment may be
diluted and mixed 1:1 with overlying water to facilitate sieving (22), see Section 6, Interferences.
1033 If the experimental design prescribes not sieving a field collected sediment, obvious large predators or
other large organisms should be removed by using forceps. If sediment is to be collected from multiple field
samples and pooled to meet technical objectives, the sediment should be thoroughly homogenized by stirring, or
with the aid of a rolling mill, feed mixer, or other suitable apparatus (see ASTM Proposed Guide for Collection,
Storage, Characterization, and Manipulation of Sediments for Toxicological Testing).
103.4 Additional samples may be taken from the same grab for other kinds of sediment analyses (see 10.1).
Qualitadve descriptions of the sediment may include color, texture, presence of macrophytes, animals, tracks, and
burrows. Monitoring the odor of sediment samples should be avoided because of hazardous volatile
contaminants (see Section 7, Safety Precautions).
103.5 The natural geochemical properties of test sediment collected from the field must be within the
tolerance limits of the test species. The limits for the test species should be determined experimentally in
advance (see 102). Controls for such factors as particle size distribution, organic carbon content, pH, etc., should
be run if the limits are exceeded in the test sediments (23).
10.4 Laboratory Spiked Sediment ~ Test sediment can also be prepared in the laboratory by manipulating
the properties of the negative control or the reference sediment. This can include adding chemicals or complex
waste mixtures (see Section 1.4) (24). The toxicity of substances either dissolved in the interstitial water or
adsorbed to sediment particles can be determined experimentally.
10.4.1 The test material(s) should be reagent grade7 or better, unless a test on formulation commercial
product (25), or technical-grade or use-grade material is specifically needed. Before a test is started, the
following should be known about the test material (a) the identity and concentration of major ingredients and
impurities, (b) water solubility in test water, (c) estimated toxicity to the test species and to humans, (d) precision
and bias of the analytical method at the planned concentration(s) of the test material, if the test concentration(s)
are to be measured, and (e) recommended handling and disposal procedures. The toxicity of the test material in
sediments may be quite different from the toxicity in water borne exposures.
10.4.2 Stock Solution(s). Test material(s) to be tested in sediment should be dissolved in a solvent to form a
stock solution that is then added to the sediment. The maximum concentration of the solvent in the sediment
should be at a concentration that does not affect the test species. The concentration and stability of the chemical
in the stock solution should be determined before beginning the test. If the chemical(s) is subject to photolysis,
the stock solution should be shielded from the light both before and during the process of mixing into the
sediment. If a solvent other than water is necessary (the preferred solvent is water), it should be one which can
7 "Reagent Chemicals, American Chemical Society Specifications," Am. Chemical Soc., Washington, DC.
For suggestions on the testing of reagents not listed by the American Chemical Society, see "Reagent
Chemicals and Standards," by Joseph Rosin, D. Van Nostrand Co., Inc, New York, NY, and the "United
States Pharmacopeia."
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be driven off (for example, evaporated) leaving only the test chemical on the sediments. Concentrations of the
chemical in the water and sediment should be monitored before the test begins.
10.43 If a solvent other than water is used, both a sediment solvent control, and a sediment negative control
or reference sediment must be included in the test. The solvent control must contain the highest concentration of
solvent present and must use solvent from the same batch used to make the stock solution (see ASTM E 729).
The same concentration of solvent should be used in all treatments.
10.43.1 Triethylene glycol is often a good organic solvetn for preparing stock solutions because of its low
toxicity to aquatic animals, low volatility, and ability to disslove many organic chemicals. Other water-miscible
organic solvents, such as methanol, ethanol or acetone may be used, but they might affect total organic carbon
levels, introduce toxicity, alter the geochemical properties of the sediment, or stimulate undesirable growths of
microoorganisms (see Section 6, Interferences). Acetone is highly volatile and might leave the system more
readily than methanol or ethanoL A surfactant should not be used in the preparation of a stock solution because
it might affect the bioavailability, form and toxicity of the test material.
10.4.4 If the concentration of solvent is not the same in all test solutions that contain test material, either (a)
a solvent test should be conducted to determine whether survival, growth, or reproduction of the test organisms is
related to the concentration of the solvent over the range used hi the toxicity test, or (b) such a solvent test
already conducted using the same overlying water and test species. If survival, growth, or reproduction is found
to be related to the concentration of solvent, a sediment toxicity test with that species in that amount of solvent is
unacceptable if any treatment contained a concentration of solvent hi that range.
10.4.4.1 If the test contains both a negative control and a solvent control, the survival, growth, or
reproduction of the organisms tested in the two controls should be compared (see ASTM E 1241). If a
statistically significant difference in either survival, growth, or reproduction is detected between the two controls,
only the solvent control may be used for meeting the acceptability of the test and as the basis for calculation of
results. The negative control might provide additional information on the general health of the organisms tested.
If no statistically significant difference is detected, the data from both controls should be used for meeting the
acceptability of the test and as the basis for calculation of results (see ASTM E 1241, Section 9.2.43).
10.4.5 Test Concentration(s) for Laboratory Spiked Sediments.
10.4.5.1 If the test is intended to allow calculation of an LC50, the test concentrations should bracket the
predicted LC50. The prediction might be based on the results of a test on the same or a similar test material on
the same or a similar species. The LC50 of a particular compound may vary depending on physical and chemical
sediment characteristics. If a useful prediction is not available, it is desirable to conduct a range-finding test hi
which the organisms are exposed to a control and three or more concentrations of the test material that differ by
a factor of tea.
10A5.2 If necessary, concentrations above aqueous solubility can be used, as indigenous organisms are at
times exposed to concentrations above solubility in the real world (see ASTM E 729).
10A53 Bulk sediment chemical concentrations might be normalized to factors other than dry weight. For
example, concentrations of non-polar organic compounds might be normalized to sediment organic carbon
content, and metals normalized to acid volatile sulfides.
10.4.5.4 In some situations (for example, regulatory) it might be necessary to only determine (a) whether a
specific concentration of test material is toxic to the test species, or (b) whether the LC50 is above or below a
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specific concentration. When there is interest in a particular concentration, it might only be necessary to test that
concentration and not to determine the LC50.
10.4.6 Addition of test material(s) to sediment may be accomplished using various methods, such as a (a)
rolling mill, (b) feed mixer, or (c) hand mixing, (see ASTM Guide for Collection, Storage, Characterizadon, and
Manipulation of Sediments for Toxicological Testing).
10.4.6.1 Modifications of the mixing techniques might be necessary to allow time for a test material to
equilibrate with the sediment If tests are repeated, mixing conditions such as duration and temperature of
mixing, and time of mixing before the test starts, should be kept constant. Care should be taken to ensure that a
test material added to sediment is thoroughly and evenly distributed within the sediment. If necessary,
subsamples of the sediment within a mixing container can be analyzed to determine degree of mixing and
homogeneity.
11. Test Organisms
11.1 Species — Whenever possible and appropriate, tests should be conducted with species listed in the
Appendices. Use of these species is encouraged to increase comparability of results. The source and type of
sediment being tested or the type of test to be implemented might dictate selection of a particular species. The
species used should be selected based on (a) availability, (b) sensitivity to a test material(s), and (c) tolerance to
ecological conditions such as temperature, grain size, and ease of handling in the laboratory. The species used
should be identified using an appropriate taxonomic key.
1L2 Age — All organisms should be as uniform as possible in age and size class. The age or size class for a
particular test species should be chosen so that sensitivity to test materials is not affected by state of maturity,
reproduction, or other intrinsic life-cycle factors (see Species Specific Appendices).
113 Source — All organisms in a test must be from the same source. Organisms may be obtained from (a)
laboratory cultures, (b) commercial, state or federal institutions, or (c) natural populations from clean areas.
Laboratory cultures of test species can provide organisms whose history, age, and quality are known. Local and
state agencies might require collecting permits.
11.4 Quality - Analysis of the test organisms for the test material(s) is desirable, as it might be present in
the environment, and other chemicals to which major exposure might have occurred.
1L5 Brood Stock ~ Brood stock should be cared for properly so as not to be unnecessarily stressed (see
Species Specific Appendices). To maintain organisms b good condition and avoid unnecessary stress, they should
not be crowded and should not be subjected to rapid changes in temperature or water quality characteristics.
11.6 Handling - Test organisms should be handled as little as possible. When handling is necessary, it
should be done as gently, carefully, and as quickly as possible. Organisms should be introduced into solutions
beneath the air-water interface. Any organisms that touch dry surfaces, are dropped, or injured during handling
should be discarded.
12. Experimental Design
12.1 Decisions concerning the various aspects of experimental design, such as the number of treatments,
number of test chambers and test organisms per treatment, and water quality characteristics, should be based on
the purpose of the test and the type of procedure that is to be used to calculate results (see Section 16,
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Calculation of Results). A test intended to allow calculation of a specific endpoint such as an LC50 should
consist of a negative control sediment, a solvent control(s), a reference sediment, and several test sediments (see
Section 10, Sediment Characterization).
122, The object of a qualitative reconnaissance survey is to identify sites of toxic conditions that warrant
further study. It is often conducted in areas where little is known about contamination patterns. To allow for
maximum spatial coverage, the survey design might include only one sample from each site. The lack of
replication usually precludes statistical comparisons, but identification of samples for further study is possible,
where survival, growth, or reproduction differ from the negative control or reference sediment. A useful
summary of field sampling design is presented by Green (26).
122.1 The object of a quantitative statistical comparison is to test for statistically significant differences in
effects (see Section 13.12, Biological Data) among negative control or reference sediments and test sediments
from several sites. The number of replicates needed per site is a function of the need for sensitivity or power.
Replicates (for example, separate samples from different grabs taken at the same site) should be taken at each
site in the survey. Separate subsamples from the same grab might be used to test for within-grab variability, or
split samples of composited sediment from one or more grabs might be used for comparisons of test procedures
(such as comparative sensitivity among test species), but these subsamples should not be considered to be true
replicates for statistical comparisons among sites.
1222 Stir locations might be distributed along a known pollution gradient, in relation to the boundary of a
disposal site, or at sites identified as being toxic in a reconnaissance survey. Comparisons can be made in both
space and time (see Calculation of Results, Section 16). In pre-dredging studies, a sampling design can be
prepared to assess the toxicity of samples representative of the project area to be dredged. Such a design should
include subsampling cores taken to the project depth.
123 Laboratory Experiments. The primary focus of the physical and experimental test design, and statistical
analysis of the data, is the experimental unit, which is defined as the smallest physical entity to which treatments
can be independently assigned (27). Because overlying water or air can not flow from one test chamber to
another the test chamber is the experimental unit (see Section 8.4, Test Chambers). As the number of test
chambers per treatment increases, the number of degrees of freedom increases, and, therefore, the width of the
confidence interval on a point estimate, such as an LC50, decreases, and the power of a significance test increases
(see Calculation of Results, Section 16). Because of factors that might affect results within test chambers and
results of the test: (a) all test chambers should be treated as similarly as possible, such as temperature and
lighting (unless these are the variables tested), and (b) each test chamber, including replicate test chambers, must
be physically treated as a separate entity. Treatments must be randomly assigned to individual test chamber
locations. Assignment of test organisms to test chambers must be randomized.
13. Procedure
13.1 Sftdimmf inffl Tgst Oiamhers - The day before the toxicity test is started (Day -1) each test sediment,
reference sediment, and negative control sediment should be mixed and a sample added to the test chambers
(4,24,28). Sediment depth in the test chamber is dependent on the experimental design and the test species (see
Species Specific Appendices and Section 6.12). Each test chamber and replicates must contain the same amount
of sediment, determined either by volume or weight.
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13.1.1 The sediment aliquot in each test chamber should be settled by smoothing with a utensil constructed
of a suitable material (see Section 8.2, Construction Materials). If beakers are used, bubbles can be removed by
either tapping the test chamber against the palm of the hand or by displacement of bubbles with the utensil.
After the sediment is placed in the test chambers, overlying water should be added. The overlying water should
be gently poured along the side of the test chamber to prevent resuspension of the sediment.
132 Stati^ Jewing ~ Overlying water should be added to the test chambers at the volume specified by the
experimental design. Watch glasses should be used to cover the test chambers and overlying water gently
aerated. Aeration can be provided to each test chamber through a 1-mL glass pipet that extends between the
beaker spout and the watch glass cover to a depth not closer than 2 cm from the sediment surface. Air should be
bubbled into the test chambers at a rate that does not cause turbulence or disturb the sediment surface. To
allow any suspended sediments to settle, the test organisms should not be introduced into the test system for 12-
24 hours. Water quality characteristics should be measured prior to the addition of the test organisms (see
Section 13.11, Overlying Water Quality Measurements).
13.2.1 Water lost to evaporation or splattering should be replaced as needed with temperature acclimated
de-ionized water or overlying water. The water quality of the overlying water hi static sediment toxitity tests
(water hardness, alkalinity, total dissolved solids, and dissolved oxygen) might be altered by the presence of
sediment (4) or by the addition of food to the test chamber (3). These changes in water quality characteristics
might influence the availability of contaminants to the test organisms (see Section 6, Interferences).
133 Flow-Through Testing — The water-delivery system should be turned on before a test is started to verify
that the system is functioning properly. The water flow to each test chamber should not differ by more than 10%
(see Section 83.1). The total volume flow per hour for continuous flow diluters should be recorded.
133.1 After the sediment has been added (Day -1), overlying water is added to the test chambers (see
Section 132 Static Testing). After aliquots are removed for water quality determinations (Day 0), overlying water
flow is started prior to the addition of the test organisms and food (4).
13.4 Duration of Test — The test begins when test organisms are first placed hi the test chambers (Day 0)
and continues for the duration specified in the experimental design for a specific test organism (see Species
Specific Appendices).
13.5 Dissolved Oxygen - The dissolved oxygen concentration in each test chamber should be measured in at
least one test chamber in each treatment (a) at the beginning and end of the test and at least weekly (if possible)
during the test, (b) whenever there is an interruption of the flow of air (static tests) or water (flow-through tests),
and (c) whenever the behavior of the test organisms indicate that the dissolved oxygen concentration might be too
low (for example, emergence from the sediment). A measured dissolved oxygen concentration should be > 40%
and < 100% saturation (E 729, Section 12.4.2).
13.6 Ovgr|y|ng Water Quality Measurements - Conductivity, hardness, pH, and alkalinity should be
measured in all treatments at the beginning and end of a short-term test, and at least weekly during a long-term
test, using appropriate ASTM standards when possible.
13.1 Temperature - Test temperature depends upon the species used (see Species Specific Appendices).
Other temperatures may be used to study the effect of temperature on survival, growth, or reproduction of test
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organisms, and contaminant related properties (for example, bioavailability). The daily mean test temperature
must be within ± 1°C of the desired temperature. The instantaneous temperature must always be within ± 3°C
of the desired temperature.
13.8 Feting — Recommended food, ration, method and frequency of feeding test organisms are contained
in Species Specific Appendices. The food used should be analyzed for the test material and other possible
contaminants. A batch of food may be used if it will support normal function. Detailed records on feeding rates
and appearance of the sediment should be made daily.
13.9 Debris — Any floating debris may be skimmed from the test chambers before test organisms are added.
This can be accomplished with a piece of fine nylon screen or other suitable material If more than 0.1 g of
floating debris is removed, an analysis should be performed to determine the amount of chemical removed from
the system (25).
13.10 TJghf — For sediment toxicity tests various Hghf;darlcnc^$ regimes can be used depending on the
species being tested (see Species Specific Appendices) and various experimental designs.
13.11 Animation - Test organisms should be acclimated if they are cultured in water different from the
overlying water or temperature (4) (see Species Specific Appendices).
13.12 Biological Data -- Effects indicating toxicity of test sediment include mortality and sublethal effects on
growth, maturation, behavior, and reproduction. Test chambers should be observed at least daily. At the end of
the exposure period, recovery of the test organisms from sediments should be accomplished following the
methods outlined for each species (see Species Specific Appendices).
13.13 Other Measurements:
13.13.1 Field Sediment. Sediment samples should be collected from the same grab for analysis of sediment
physical and chemical characterizations. A separate sample for benthic fauna! analyses may be desirable (see
ASTM D 4387).
O.13.2 Laboratory Spiked Sediments. At the beginning and at the end of the experiment, measurement of
the concentration of the test material(s) in both stock solutions and sediment, is desirable. To monitor changes
in sediment or interstitial water chemistry during the course of the experiment, separate sediment chemistry
chambers should be set up and sampled at the start and end of the experiment. It is not necessary to add test
organisms to these chambers at the beginning of the test, but for later sampling, test organisms should be added
after the initial sample is taken.
13.13.2,1 Concentration of test material(s) in overlying water, interstitial water and sediment should be
measured at several concentrations and as often as practical during the test If possible, the concentration of the
test material in overlying water, interstitial water and sediments should be measured at the start and end of the
test. Measurement of test material(s) degradation products might also be desirable.
13.13.22 Measurement of test material(s) concentration in water can be accomplished by pipeting water
samples from a point midway between top, bottom and sides of the test chamber. Overlying water samples
should not contain any surface scum, any material from the sides of the test chamber, or any sediment
13.1323 Measurement of test material(s) concentration in sediment at the end of a test can be taken by
siphoning the overlying water without disturbing the surface of the sediment, then removing appropriate aliquots
of the sediment for chemical analysis.
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13.132.4 Interstitial water can be sampled by using the water that (a) comes to the surface in a mixing
apparatus, (b) is on the surface of the sediment after it settles, (c) is separated from the sediment particles by
centrifuging a sediment sample, (d) is filtered through an apparatus to extract interstitial water, (e) has been
pressed out of the sediment, or (f) by using an interstitial water sampler. Care should be taken to ensure that
contaminants do not transform, degrade, or volatilize during the interstitial water sample preparation (see ASTM
Guide for Collection, Storage, Characterization, and Manipulation of Sediments for Toxicological Testing).
14. Analytical Methodology
14.1 Chemical and physical data should be obtained using appropriate ASTM standards whenever possible.
For those measurements for which ASTM standards do not exist or are not sensitive enough, methods should be
obtained from other reliable sources (29).
14.2 Concentrations should be measured for (a) contaminants in bulk sediment, (b) test material(s) in the
interstitial water, (c) test material(s) in the overlying water, and (d) test material(s) in the stock solution. In
addition, measurement of either the apparent dissolved or undissolved substances of the test material(s) is
desirable. The apparent dissolved material is defined and determined as that which passes through a 0.45 /*m
membrane filter.
14.2.1 If samples of overlying water from test chambers, stock solutions, test sediment or interstitial water
are not to be analyzed immediately, they should be handled and stored appropriately (30) (see Section 10,
Sediments).
143 Methods used to analyze food or test organisms should be obtained from appropriate sources (31).
14.4 The precision and bias of each analytical method used should be determined hi an appropriate matrix:
that is, sediment, water, tissue. When appropriate, reagent blanks, recoveries, and standards should be included
when samples are analyzed.
15. Acceptability of Test
L5.1 A sediment toxicity test should be considered unacceptable if one or more of the following occurred,
except, for example, if temperature was measured numerous times, a deviation of more than 3°C (see 13.6,
Temperature) in any one measurement might be inconsequential. However, if temperature was measured only a
minimal number of tunes, one deviation of more than 3°C might indicate that more deviations would have been
found if temperature had been measured more often.
15.1.1 All test chambers (and compartments) were not identical (Section 8.4.1, 123).
15.12 The overlying water was not acceptable to the test organisms (Section 9.1).
15.13 Test organisms were not acclimated to the approriapte overlying water or temperature if they are
cultured in water different from the overlying water or temperature.
15.1.4 The natural geochemical properties of test sediment collected from the field was not within the
tolerance limits of the test species (Section 103.5).
15.1.5 Appropriate negative and solvent controls, or reference sediment, were not included in the test
(Section 10.43).
15.1.6 The concentration of solvent in the range used affected survival, growth, or reproduction of the test
organisms (Section 10.4.4).
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15.1.7 All animals in the test population were not obtained from the same source, were not all of the same
species, or were not of acceptable quality (Section 113).
L5.L8 Treatments were not randomly assigned to individual test chamber locations and the individual test
organisms were not impartially or randomly assigned to test chambers or compartments (Section 123).
15.1.9 Each test chamber must contain the same amount 01 sediment, determined either by volume or
weight.
15.1.10 Temperature, dissolved oxygen, and concentration of test material were not measured, or within the
acceptable range (Section 13.7 and Species Specific Appendices).
15.1.11 The negative control or reference sediment organisms did not survive, grow or reproduce as required
for the test species (see Species Spc opendices).
L5.1.12 Average survival in any negative control chamber is less than acceptable limits (see Sp -s Specific
Appendices).
16. Calculation of Results
16.1 The calculation procedure(s) ard interpretation of the results should be appropriate to the experimental
design. Procedures used to calculate resuAs of toricity tests can be divided into two categories: those that test
hypotheses and those that pro ..- point estimates. No procedure snould be used without careful consideration of
(a) the advantages and disadvantages of various alternative procedures, and (b) appropriate preliminary tests,
such as those for outliers and for heterogeneity.
16.2 For each set of data the LC50 or EC50 and its 95% confidence limits should be calculated (when
anoropriate) on the basis of (a) the measured initial concentrations of test material, if available, or the calculated
initial concentrations for . tatic tests, and (b) the average measured concentrations of test material, if available, or
the calculated average concentrations for flow-through tests. If other LC or ECs are calculated, their 95%
confidence limits should also be calculated (see ASTM E "29).
163 Most toricity tests produce quanta! data, that is, counts of the number of responses in two mutually
exclusive categories, such as alive or dead. A variety of methods (32) can be used to calculate an LC50 or EC50
and 95% confidence limits from a set of quantal data that is binomially distributed and contains C.VG or more
concentrations at which the percent dead or effected is between zero and 100, but the most widely used are the
probit, moving average, Spearman-Karber and Litchfield-Wilcoxon methods. The method used should
appropriately take into account the number of test organisms per chamber. The binomial test can also be used
to obtain statistically sound information about the LC50 or EC50 even when less than two concentrations kill or
affect between zero and 100 percent. The binomial test provides a range within which the LC50 or EC50 should
lie.
16.4 When samples from field sites are independently replicated, the site effects can be statistically compared
by t-tests, analysis of variance (ANOVA) or regression type analysis. Analysis of variance is used to determine
whether any of the observed differ -.aces among the concentrations (or samples) are statistically significant. This
is a test of the null hypothesis tk.. no differences exist in the effects at all of the concentrations (or samples) anu
at the control If the F-test is not statistically signifiranr (P>0.05), it can be concludeu that the effects observed
in the test material treatments (or field sites) were not large enough to be detected as statistically significant by
the experimental design and hypothesis test used. Non-rejection does not mean that the null hypothesis is true.
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The NOEC based on this end point is then taken to be the highest test concentration tested (3334). The amount
of effect that occurred at this concentration should be considered.
16.4.1 All exposure concentration effects (or field sites) can be compared with the control effects by using
mean separation techniques such as those explained by Chew (35) orthagonal contrasts, Fisher's methods,
Dunnett's procedure or Williams' method. The lowest concentration for which the difference in observed effect
exceeds the statistical significant difference is defined as the LOEC for that end point. The highest concentration
for which the difference in effect is not greater than the statistical significant difference is defined as the NOEC
for that end point (33).
17. Documentation
17.1 The record of the results of an acceptable sediment toricity test should include the following
information either directly or by reference to available documents.
17.1.1 Name of test and investigator(s), name and location of laboratory, and dates of start and end of test.
17.1.2 Source of negative control, reference or test sediment, method for collection, handling, shipping,
storage and disposal of sediment.
17.13 Source of test material, lot number if applicable, composition (identities and concentrations of major
ingredients and impurities if known), known chemical and physical properties, and the identity and
concentration(s) of any solvent used.
17.1.4 Source of overlying water, its chemical characteristics, and a description of any pretreatment, and
results of any demonstration of the ability of a species to survive, grow or reproduce in the water.
17.1.5 Source, history and age of test organisms; source, history and age of brood stock, culture procedures;
and source and date of collection of the test organisms, scientific name, name of person who identified the
organisms and the taxonomic key used, age, life-stage, means and ranges of weight and lengths, observed diseases
or unusual appearance, treatments, holding and acclimation procedures.
17.1.6 Source and composition of food, concentrations of test material and other contaminants, procedure
used to prepare food, feeding methods, frequency and radon.
17.1.7 Description of the experimental design and test chambers (and compartments), the depth and volume
of sediment and overlying water in the chambers, lighting, number test chambers and number of test organisms
per treatment, date and time test starts and ends, temperature measurements, dissolved oxygen concentration (as
percent saturation) and any aeration used prior to initiating a test and during the conduct of a test.
17.1.8 Methods used for, and results (with standard deviations or confidence limits) of, physical and chemical
analyses of sediment.
17.L9 Definition(s) of the effects used to calculate LC50 or ECSOs, biological endpoints for tests, and a
summary of general observations of other effects.
17.1.10 A table of the biological data for each test chamber for each treatment including the control(s) in
sufficient detail to allow independent statistical analysis.
17.1.11 Methods used for, and results of, statistical analyses of data.
17.1.12 Summary of general observations on other effects or symptoms.
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17.1.13 Anything unusual about the test, any deviation from these procedures, and any other relevant
information.
17.1.14 Published reports should contain enough information to dearly identify the methodology used and
the quality of the results.
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ANNEX XI. Hvalella azteca
Xl.l Significance - Hvalella azteca (Saussure), Amphipoda, has many desirable characteristics of a test
species: short generation time, easily collected from natural sources or cultured in the laboratory in large
numbers, and data on survival, growth, and reproduction can be obtained in toxicity tests (36). Landrnm and
Scavia (37), Nebeker et al. (22), and Ingersoll and Nelson (4) have successfully used H. azteca in sediment
toxicity testing and have shown it to be a sensitive indicator of the presence of contaminants associated with
sediments. Ingersoll and Nelson (4) report _H. azteca to have a wide tolerance of sediment grain size. Sediment
ranging from >90% silt- and clay-size particles to 100% sand-size particles did not reduce survival or growth in
the laboratory.
X12 Life History and Life-Cvcle - The life-cycle of H. azteca can be divided into three distinct stages
according to Cooper (36): (1) an immature stage, consisting of the first 5 instars; (2) a juvenile stage, including
instars 6 and 7; and (3) an adult stage, the 8th instar and older. The potential number of adult instars is large
and growth is indeterminate such that old adults can be much larger than younger adults (38). DeMarch (39)
indicates that juvenile H. azteca. can complete a life-cycle in 27 days or longer depending on temperature.
X 1.2.1 _H- azteca is an epibenthic detritivore and will burrow in the sediment surface, and Hargrave (40) has
demonstrated in laboratory experiments that H. azteca digests bacteria and algae from ingested sediment
panicles (< 65 /jm), further illustrating sediment interactions bv H. azteca.
X1.12 Sexual dimorphism occurs in H. azteca. the adult male is larger than females and has larger second
gnathopods (41).
X1.23 DeMarch (41) indicates that the number of young produced per adult female is optimum at
temperatures of 26-28 °C. Whereas, Cooper (36) and Strong (38) report that maximum brood size is more
dependent on the size of the adult amphipods than on temperature.
XL3 plaining Test Organisms . The following culture procedures are adapted from deMarch (41),
Nebeker et aL (22), and Ingersoll and Nelson (4). H. azteca can be reared in 10- or 20-L aquaria under flowing
water conditions with a 16:8 hour light:darkness photoperiod at 20_+.2°C, and about 500 foot-candles (5382 lux).
For static cultures, the water should be gently aerated and about 25-30 percent of the water volume should be
replaced weekly. In flow-through cultures, water delivery can be at a low rate (100 mL/min) (4).
XI J.I .H,. azteca can be cultured with a variety of foods. Dried maple, alder, birch or poplar leaves,
prcsoaked for several days and tannins flushed out with water, then can be added weekly as the primary substrate
and food. Rabbit pellets 8, ground cereal leaves 9, fish food flakes 10, frozen or newly hatched brine shrimp or
heat-killed vounp Daphnia can be used to feed H. azteca. In addition, Strong (38) demonstrated success in
* Purina Rabbit Chow, Purina Mills, Inc, 1401 Hanley, St. Louis, MO 63144.
9 Ccrophyl, Sigma Chemical Company, P.O. Box 14508, St. Louis, MO 63178.
10 TetraMin Fish Food Flakes, TetraWerke, Dr. rer. naL Ulrich Baensch GmbH, D-4520 Melle 1,
W. Germany.
21
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culturing H. azteca yielding the best survivorship and consistently the largest dutches by feeding the amphipods
filamentous green algae (QcdflgonJUTTH cardiacum^ and homogenized rotting spinach ad libitum.
XL3.2 To dean the culture tanks or reduce populations of animals, half of the leaf substrate containing a
portion of the animals should be transferred to a sorting tray, discarding the remainder of the old contents and
returning the leaf substrate and animals to the chamber. The number of amphipods should be reduced
periodically as the population expands rapidly.
X1.4 Collection - _H. azteca can be found in permanent lakes, ponds and streams throughout the entire
American continent (41,42). Methods used by Landrnm and Scavia (37) indicate that the amphipods can be
collected from a natural freshwater source. Pennak (42) suggests using a dip-net to collect aquatic vegetation and
bottom debris containing amphipods. Sites with stony bottoms might require collecting with forceps or the use a
small aquarium net Live specimens can be maintained in aquaria if they are well supplied with aquatic
vegetation (42). Collection procedures for K azteca, by deMarch (41) indicate that rinsing aquatic vegetation is
effective if a 200-550 fim mesh net is used to catch the amphipods. Up to 200 amphipods can be transported in a
large plastic bag containing 1 L of water from the collection site, with the remainder of the bag filled with air or
oxygen and then placed into a cooler (41). For verification and accurate identification of field collected H.
azteca. it is important that mature males and females be used (42).
X1.5 Brood Stock - Brood stock can be obtained from the wild, another laboratory or a commercial source.
H. azteca brought into the laboratory should be acclimated to the culture water by gradually changing the water
in the culture chamber from the water in which they were transported to 100% culture water. _H. azteca should
be acclimated to the culture temperature by changing the water temperature at a rate not to exceed 2°C within
24 h, until the desired temperature is reached (41). Brood stock should be cultured so they are not unnecessarily
stressed. To maintain H. azteca in good condition and avoid unnecessary stress, crowding andjpapid changes in
temperature and water quality characteristics should be avoided.
X1.6 Hailing - JH[. azteca should be handled as little as possible. When handling is necessary, it should be
done as gently, carefully, and quickly as possible, so that the amphipods are not unnecessarily stressed.
Amphipods should be introduced into solutions beneath the air-water interface (4). Any .H. azteca that touch dry
surfaces, are dropped, or injured during handling should be discarded. Removing animals from sieves may form
air bubbles on body surfaces causing animals to float on the water surface. Any "floaters" should be gently placed
into the water column using a probe. If the animals continue to float they should be removed and discarded.
XL7 Age - Tests with H. azteca should be started with juvenile organisms, (second or third instar) about 2-
3 mm in length (4,22). To obtain fl. azteca for testing, amphipods should be separated from the leaf material by
scooping up the leaves with dinging amphipods, and placing the leaves on a 5-10 mm mesh screen, which is
placed over a collecting pan containing 2 cm of culture water. Culture water should be sprinkled on the leaves
while turning and separating the leaves. Mixed age H. azteca should be washed from the leaves and drop
through the screen into a collecting pan (22). To separate the juvenile amphipods from the larger adults a sieve
stack (US. Standard) #30 (600 fjaa), #40 (425 paa), and a #60 (250 urn) can be used (4). Culture water should
be rinsed through the sieves and juvenile animals retained by the #60 sieve are washed into a collecting pan
while the larger animals in the top sieves (#30 and #40) are returned to the culture. The juvenile amphipods arc
then placed in 1-L beakers containing culture water (about 200 amphipods/beaker) and kept in the dark at the
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
temperature of the culture with gentle aeration. JJ. azteca can be isolated in the 1-L beakers up to 24 hours
prior to the start of the sediment toxicity test.
Xl.7.1 Borgmann (43) recommends collecting uniform aged young (< 1 week old) for experimental
purposes using Zf-L jars containing about 1 L of culture water and 5-25 adult JH. azteca. The jars are placed in
an incubator at 16 to 8 hour light to darkness photoperiod, about 500 foot-candles (5382 lux). Each jar contains
pieces of pre-soaked (in culture water) cotton gauze as a substrate. Once a week the animals should be removed
from the gauze and collected by filtration through a 275 /im nylon mesh screen, then rinsed into petri dishes
where the young and adults are sorted. Fresh culture water and food should be placed in the jars and the adults
returned. Each jar should receive 0.02 g of fish food flakes 10 or more if required by larger animals.
X1.8 Ary1imatifln - If amphipods are cultured in water different from the overlying water or temperature, an
acclimation process is necessary. The water acclimation process used by Ingersoll and Nelson (4) is to first place
animals for 2 h in a 50:50 mixture of culture water to overlying water, then for 2 h in a 25:75 mixture of culture
water to overlying water, followed by a transfer into 100% overlying water. At this stage the amphipods are
considered acclimated to the overlying water and are ready for immediate use. JH. azteca can then be randomly
selected from the acclimation water with a pipette and placed into counting beakers (for example, 30-mL) that
can be floated hi the test chambers before the amphipods are introduced into the exposure system (4).
X1.9 Toxicitv Test Specifications
Xl.9.1 Experimental Design - Decisions concerning the various aspects of experimental design, such as the
number of treatments, number of test chambers and amphipods per treatment, and water quality characteristics,
should be based on the purpose of the test and the procedure used to calculate results. Nebeker et al. (22)
recommend two or more replicate 20-L aquaria per treatment with 100 juvenile H. azteca placed in each
aquarium. Ingersoll and Nelson (4) recommend four replicate 1-L beakers per treatment, with 20 H- azteca per
replicate, for a total of 80 amphipods per treatment. Duration of the test can range from a _<.10 day short-term
test to a long-term test > 10 days and continuing up to 30 days (4,22). The number of young and adult survival
(4,22), growth, and development (4) can be used as the biological endpoints. A test duration up to 30 days can
add potential reproductive capacity as another biological endpoint, measuring effects on reproductive behavior,
appearance of secondary sex characteristics, egg production, and number of young produced. Tests with_H.
azteca have been conducted at 20°C (4,22) and from 21-25°C (37), photoperiod 16 to 8 hour light to darkness,
about 50 foot-candles (538 lux) (4).
XL92 Static and Flnw-thrnugh Tesr* - Ingersoll and Nelson (4) and Nebeker et al. (22) recommend using
borosilicate glass 1-L beakers to expose the JH- azteca to the test material These exposure chambers contain
about 800 mL overlying water and 200 mL (2 cm) test sediment, in both the static and flow-through water
systems. For the static tests cover watch glasses may be used to fit over the top, such that an aeration dp fits
through the beaker pour spout and the cover (4). Nebeker et aL (22) suggest for the static long-term test, using
20-L aquaria with 2 - 3 cm of test sediment on the bottom overlaid with 15 cm water. For flow-through testing,
Ingersoll and Nelson (4) suggest using a 4 x 13 cm notch cut in the lip of the 1-L beaker. The notch should be
covered with 033 mm U.S. Standard sieve size #50 screen, either made of stainless steel or polyethylene, using a
silicone adhesive to attach the screen to the beaker.
23
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X1.93 Initiation of a Test - Sediments should be homogenized and placed in the test chambers on the day
prior to the addition of the test organisms (Day -1). Test chambers should be covered and overlying water
aerated (4) or unaerated overnight but aerated for 30 minutes before _H. azteca are added (22). The test begins
when the juvenile HL azteca are introduced to the test chambers (Day 0). It is recommended that flow-through
and static tests might need to be started on different days to assure that sufficient time is available to complete
all tasks. Test chambers should be inspected <2 hours after amphipods are introduced to insure that animals are
not trapped hi the surface tension of the water (4). These "floaters* might not survive well and should be
replaced with new animals (see X1.6).
Xl.9.4 Feeding - Ingersoll and Nelson (4) recommend rabbit pellets * to be used as a food for U. azteca in
short and long-term sediment toxicity tests, Nebeker et aL (22) suggest feeding rabbit pellets 8 in a 28 day test.
The pellets should be ground and dispersed hi deionized water. A flurocarbon plastic stir bar and a magnetic stir
plate should be used to homogeneously resuspend the rabbit pellets ' when aliquots are removed for feeding. If
food collects on the sediment, a fungal or bacterial growth might start on the surface of the sediment, in which
case feeding should be suspended for one or more days. A drop in dissolved oxygen to 40% saturation might
indicate that all of the food added in the water is not being consumed such that feeding might be suspended for
the amount of time necessary to increase the dissolved oxygen concentration (4).
Xl.9.4.1 In static tests Nebeker et aL (22) suggest a feeding regime twice weekly of 200 mg (05 mL dry
volume) rabbit pellets * mixed in 100 mL distilled water for 100 juvenile H. azteca in a 20-L aquarium. Nelson
and Ingersoll (4) recommend feeding H. azteca three times weekly 14 mg rabbit pellets ' per feeding for 20
young amphipods in a 1-L beaker. Lower feeding levels for flow-through and static tests may be used for _H.
azteca: three times weekly 6 mg rabbit pellets per 8 feeding for the first week of the test, and 12 mg per feeding
for the following weeks.
Xl.9.4.2 For flow-through testing, prior to starting a test, 20 mg rabbit pellets ' should be added to each test
chamber, and three times a week each test chamber should be fed 20 mg per feeding for 20 young JH. azteca
during the exposure (4).
X1.10 Biological Data - During the conduct of the test, observations should be made to assess behavior (for
example, 'floaters', sediment avoidance) and reproductive activities (for example, amplexus). At the end of the
test the H. azteca must be removed from the test chambers for survival (4,22), observable behavior, any
noticeable reproduction (for example, amplexus, gravid females, young present) and growth (4). According to
Ingersoll and Nelson (4) without material above the sediment surface, such as the leaves used in culturing, H.
azteca burrow in the top 1 cm sediment surface or are found swimming m the water column. Many of the
surviving amphipods can be pipeted from the water column before sieving the sediments. At the end of the test
the sediment should be screened using a #35 (500 /un) U.S. Standard size sieve (22). Ingersoll and Nelson (4)
recommend using a #50 (300 /im) U.S. Standard size screen cup first by swirling the overlying water to suspend
the upper 1 cm of sediment and pouring that slurry into the cup. Next, a stack of sieves #25 and #40 U.S.
Standard ««»•- should be used to sieve the bulk sediment in order to collect and count the live animals remaining
hi the sediment. The H. fl^eca, arc rinsed from the screens into collecting pans and pipeted from the rinse water
(4). It might be difficult to recover voung H. azteca due to then- small size. Material retained in the collecting
pans may be preserved in a sugar formalin mixture for examination at a later date (4). The preserved material
may be inspected using a low power binocular microscope to search for .H. azteca missed the last day of the test.
24
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
Xl.10.1 For quantifying growth. H. azteca body length (±0.01 mm) should be measured from the base of
the first antenna to the tip of the third uropod along the curve of the dorsal surface (4). In addition, wet and dry
weight measurements have been used to estimate growth for _H. j}zt££a. (37).
Xl.10.2 A H. azteca sediment toxicity test, independent of duration, is unacceptable if the average survival in
any negative control chamber is less than 80% (see Section 15, Acceptability of Test).
25
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ANNEX X2. Chironomus tentans
X2.1 S'gTlif'CTnyT - (Tilfonoroug t?"tfln5 Fabricius (Diptera: Chironomidae) has been used in sediment
toxicity tests because it is a fairly large midge with a short generation time, is easily cultured in the laboratory,
and the larvae have direct contact with the sediment by burrowing into sediment to build a case. £. tentans has
been successfully used in sediment toxicity testing and is sensitive to many contaminants associated with
sediments (22,25,44,45,46). The members of the genus are important in the diet of young and adult fish and
surface feeding ducks (47).
X22, Life History and Life-Cvcle - The classification of holometabolous insects, such as £. tentans. presents
special difficulties because each life-stage often has different ecological requirements. Further detailed studies at
the species level are needed to better understand the various physical, chemical, and biological factors that
interact to produce a suitable environment for larval development (48). £. tentans has a holarctic distribution
and is locally common in the mid-continental areas of North America (47,49,50). Sadler (51) describes the
general biology of£. iejUans.. The larval stages often inhabit eutrophic lakes and ponds. Qualitative observations
indicate larvae occur most frequently in fine sediment and detritus; however larvae reportedly inhabit sediments
with particles ranging from <0.15 mm to 2.0 mm (52). Chironomid larvae usually penetrate a few centimeters
into sediment In both lotic and lentic habitats with soft bottoms, about 95% of the chironomid larvae occur in
the upper 10 cm of substrate, very few larvae are found below 40 cm (48). Larvae are generally not found when
hydrogen sulfide is greater than 03 mg/L (52). Larvae of C. tentans are found in the field at a temperature
range between 0°C to 35°C, pH range between 7 to 10, conductivity range between 100-4000 /iS cm"1, sediment
organic carbon range between 2 and 15 percent, and at dissolved oxygen concentrations as low as 1 mg/L
(47,52,53). Sadler (51) reported that £. tentans will eat essentially any material of appropriate size.
X2^.1 The biology of£.l£fllaas facilitates laboratory culture since larvae are tolerant of a wide spectrum of
conditions and adults mate even when confined (47). The life-cycle of C. tentans can be divided into three
distinct stages: (1) a larval stage, consisting of the 4 instars; (2) a pupal stage, and (3) an adult stage. Midge egg
masses hatch in 2 or 3 days after deposition in water at 19-22°C. Larval growth occurs in four instars of about
one week each. Under optimal conditions larvae will pupate and emerge as adults after 24-28 days at 20° C.
Adults emerge from pupal cases over a period lasting several days. Males are easily distinguished from females
because males have large, plumose antennae and a much thinner abdomen with visible genitalia. Mating
behavior has been described by Sadler (51) and others (54).
X23 Obtaining TffSt Qr83I"Sm5 • The following is a description of culturing procedures adapted from
Adams et at (25), Nebeker et aL (22) and others (47,54). These procedures should not be considered definitive.
What works in one laboratory sometimes works poorly in another laboratory. £. tentans can be reared in
aquaria in static or flowing water with a 16 to 8 hour light to darkness photoperiod at 20-23° C, at about 50 foot-
candles (538 lux). For static cultures the water should be gently aerated and about 25-30 percent of the water
volume should be replaced weekly. Cultures should be maintained in an isolated area or room free of
contamination and excessive disturbances. Adams et aL (25) recommends rearing midges in glass aquaria filled
with water to a depth of 45 cm covered with nylon screen. The size of the aquaria may vary from a minimum of
3 L to a maximum of 19 L depending on the need for animals.
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
X23.1 Chironomus fentans require a substrate in which to construct a case. Shredded paper towels have
been found to be well suited for this purpose. Strips cut from Scott" or NibrocR brown paper towels should be
soaked overnight in acetone to remove impurides and are then rinsed in three changes of culture water until the
acetone is removed. A kitchen blender should be used to shred the rinsed towels into a pulp. Care must be
taken to avoid over blending and possibly shortening the wood fibers in the pulp. The pulp should be rinsed
twice with culture water to remove extremely small fibers and refrigerated until needed. The paper toweling pulp
should be placed into the water of a culture chamber to a depth of 3 on. One gram of dry fish food flakes '°
should be mixed in 10 mL of culture water with a kitchen blender and refrigerated. This suspension should be
fed twice daily to the cultures for optimum growth. The amount given depends on the number and size of the
larvae. If after feeding the culture water does not clear in 3 to 4 hours, the feeding level should be reduced.
Overfeeding will lead to the growth of fungus in the aquaria and will necessitate more frequent water changes.
Therefore, new cultures should receive 0.5 mL or less of this suspension per feeding. Nebeker et al. (22) suggest
supplementing the fish food flakes 10 diet with ground cereal leaves 3.
X2.4 Brood Stock - Brood stock can be obtained from the wild, laboratory or a commercial source. When
midges are brought into the laboratory, they should be acclimated to the culture water by gradually changing the
water in the culture chamber from the water in which they were transported to 100% culture water. Midges
should be acclimated to the test temperature by rhanging the water temperature at a rate not to exceed 2°C
within 24 h, until the desired temperature is reached. Brood stock should be cultured so they are not
unnecessarily stressed. To maintain midges in good health and avoid unnecessary stress, crowding and rapid
changes in temperature and water quality characteristics should be avoided.
X2^ Age - Test with £. tentans can be started with second instar larvae according to Wentsel et al., (44),
Adams et al. (25), Nebeker et al. (22) and Giesy (45). Tests started with first instar £. tentans larvae have met
with limited success (22). Twelve to 16 days before a test is begun, at least 3 freshly laid midge egg cases should
be placed in a dean 20x40 cm glass or enameled rearing pan filled with water to a depth of 3 cm. Egg cases
should be isolated by aspirating adults into a 250-mL Erlenmeyer flask in the morning. In late afternoon, about
20 mL of culture water should be added to the flask. Egg cases are deposited overnight and first instar larvae
begin to hatch after about 3 days at 20°C. No substrate is added to the pan before hatching. Fish food flakes 10
should be added at a rate of 50 mg/day suspended in water. Fresh water should be added as needed to make up
for evaporation. The larvae in the rearing pans are presumed to be 2nd instars on the 12th day from the time
the eggs were laid (10 day old larvae). Most larvae will remain as 2nd instars through the 16th day (14 day old
larvae). Larvae _>.16 days old should not be used to start a test. To maintain a supply of 2nd instar larvae for
active toxicity testing, a rearing pan should be started every 4 days. Each pan can be expected to produce at least
enough 2nd instar larvae for one sediment toxicity test
X2.6 Handling . Midges should be handled as little as possible. When handling is necessary, it should be
done as gently, carefully, and quickly, so that the midges are not unnecessarily stressed. Larvae should be
transferred with a 7-mm inner diameter glass pipet. Midges should be introduced into solutions beneath the air-
water interface. Any midges that touch dry surfaces, are dropped, or injured during handling should be
discarded.
27
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X2.7 ATT*iTnafifln - If the midges are cultured in water different from the overlying water or temperature, an
acclimation process is necessary. The water acclimation process used by Ingenoll and Nelson (4) is to first place
animals for 2 h in a 50:50 mixture of culture water to overlying water, then for 2 h in a 25:75 mixture of culture
water to overlying water, followed by a transfer into 100% overlying water. At this stage the midges are
considered acclimated to the overlying water and are ready for immediate use. Midges should be randomly
selected from the acclimation water with a pipette and placed into counting beakers, for example 30-mL, that can
be floated in the test chambers before the midges are introduced into the exposure system (4).
X2J& Toxicirv Test Specifications
X2JJ.1 Experiment' Dgfi'gP - Decisions concerning the various aspects of experimental design, such as the
number of treatments, number of test chambers and midges per treatment, and water quality characteristics,
should be based on the purpose of the test and the type of procedure that is to be used to calculate results.
Tests with£.i£alaflfi have been conducted at 20-23°C (22,25,44). Cooler test temperature may reduce the
growth of fungus on the sediment surface. Duration of the test can range from a _<.10 day test to > 10 days and
continuing up to 25 days (22,25,44,45). Larval survival, growth, or adult emergence can be monitored as
biological endpoints.
X2JJ.2 Static and Flow-through tests - Wentsel et al. (44) recommend using 20 £. tentans in each 2-L
exposure beaker containing 2 cm of sediment and 1-5 L of overlying water in static testing. Adams et aL (25) use
3-L aquaria constructed of glass and silicone rubber for either static or flow-through testing. These test chambers
measure 20.5 x Ii5 x 14.5 cm with a 123 x 44.5 cm piece of fine mesh stainless steel screen positioned on the
upper end of one side. This overflow screen prevents the escape of larvae and maintains an overlying water
volume of 2 L with 100 g of test sediment and 25 £. isstans. larvae per chamber. Nebeker et al. (22) recommend
20-L aquaria with 100 £. tentans larvae and 2 to 3 cm of test sediment on the bottom with 15 cm of overlying
water in static tests. If less sediment is available for testing, 4-L glass jars can be used, but proportionally fewer
animals and less food should be used. Adams et aL (25) and Giesy et aL (45) also describe a method to expose
midges individually to contaminated sediment in static tests. Up to 15 £. tentans are placed in separate 50-mL
plastic centrifuge tubes. Each tube contains one midge, 7.5 g of sediment and 47 mL of water. For 24 hours
after hatching, first instar midge larvae are often planktonic (55). If flow-through tests are started with first instar
£. tentans larvae, water flow into the test chambers should not be started for at least 24 hours after larvae are
added. This will allow time for larvae to settle onto the sediment surface.
X2JJ3 Initiation of a Test - Sediments should be homogenized and placed in the test chambers on the day
before addition of test organisms (Day -1). Test chambers should be covered and overlying water aerated
overnight The test begins when midges are introduced to the test chambers (Day 0). Larvae must be collected
from at least three separate egg cases to start a sediment toxicity test It is recommended that flow-through and
static tests might need to be started on different days to assure that sufficient time is available to complete all
tasks. Test chambers should be inspected <2 hours after midges are introduced to insure that animals are not
trapped in the surface tension of the water (4). These, "floaters" do not survive well and should be replaced with
healthy animals.
X2&4 Feeding - Adams et aL (25) recommend feeding animals in flow-through or static tests 50 mg fish
food flakes 10 (dry weight, administered in a 0.5 mL suspension) daily to each 3-L test chamber containing 25
larvae. Nebeker et aL (22) suggest feeding animals in static tests a food mixture of 600 mg ground cereal leaves 9
28
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Nelson, Coyie and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
(1.5 mL dry volume) and 100 mg (03 mL dry volume) of finely crushed fish food flakes 10 in water and feeding
this amount of food to the 100 £. T?"tan? larvae in each 20-L test chamber at the start of the test (Day 0) and on
Day 8. On day 14 they should be fed 800 mg (2.0 mL) ground cereal leaves 9 and 100 mg (03 mL) fish food
flakes 10, and on day 18 they should be fed 1,000 mg (2J mL) ground cereal leaves 9 and 100 mg (03 mL) fish
food flakes 1C. Giesy et aL (45) recommend feeding a 0.1 mL suspension of 0.06 g/mL goldfish food " daily to
each individual midge in each centrifuge tube. If food collects on the sediment, a fungal or bacterial growth
might start on the surface of the sediment, in which case feeding may be suspended for one or more days. A
drop in dissolved oxygen to 40% saturation might indicate that all of the food added in the water is not being
consumed such that feeding should be suspended for the amount of time necessary to increase the dissolved
oxygen concentration.
X2.8.5 Biological Data - Several endpoints can be monitored in midge sediment toxicity tests. During the
test, emergence of larvae from the test sediment can be monitored. Additionally, data on larval survival, growth,
and adult emergence can be obtained.
X2.8 .5.1 Larval survival and growth can be assessed by ending the tests on Day 10 to Day 14 when larvae
have readied the 3rd or 4th instar (22,25,45). At this time, larvae can be removed from sediment using a #35
(500 ftm) U.S. Standard size sieve (4). The midges can be rinsed from the sieve into collecting pans and pipeted
from the rinse water. Growth determinations using dry weight (dried at 60° C to a constant weight) is preferable
to length. Growth can also be estimated by measuring head capsule width, and also be used to determine instar
development.
X2.8.5.2 Nebeker et al. (22) suggest conducting adult £. tentans emergence sediment toxicity tests for 25
days when tests are started with second instar larvae. The adult emergence exposure chambers are covered by
screen to retain emerging adults. The adult £. tentans should begin emerging after 20 days; the test should be
continued for at least 5 days to count all the adults emerging and monitor delayed development. A small vacuum
pump with a 10-mm diameter plastic line running through an Erlenmeyer flask trap is used to collect adults and
make daily count of adults emerging. The screen cover is slowly lifted off the container and the adults are
vacuumed from the screen and inside walls of the container. Percent adult emergence is generally less than 60%
in these tests. Endpoints calculated in these adult emergence tests can include (1) percent emergence, (2) mean
emergence time, or (3) day to first emergence. Egg hatching studies may also be conducted by covering the test
chambers and confining the adults. Adults will emerge and lay eggs in these chambers. These egg masses can
then be used to estimate effects of exposure on either the number of eggs produced or hatched.
X2.8.53 A r. tqntans sediment toxicity test, independent of test duration, is unacceptable if the average
survival in any negative control chamber is less than 70% (see Section 15, Acceptability of Test). (Note: a low
percent emergence of adults might not be the result of low survival; larvae or pupae might not have completed
development).
11 TetraFin Goldfish Food, TetraWerke, Dr. rer. nat. Ulrich Baensch GmbH, D-4520 Melle 1, W.
Germany.
29
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ANNEX X3. Chironomus riparius
X3.1 fijgnjfjranc-fi - CMrQTOPiys, riparius Meigen (Diptera: Chironomidae) has been used in sediment toxicity
tests because it is a fairly large midge, has a short generation time, is easily cultured in the laboratory, and the
larvae have direct contact with the sediment by burrowing into the sediment to build a case. £. riparius has been
successfully used in sediment toxir.ity testing and is sensitive to many contaminants associated with sedi*- ints
(4,56,57,58). The members of the genus are important in the diet of young and adult fish and surface . iding
ducks (47).
X3.2 Life History and Life-Cvcle - The classification of holometabolous insects, such as £. ripar:-. . presents
special difficulties because each life-stage often has different ecological requiremer. . Further detailed studies at
the species level are needed to better understand the various physical, chemical, and biological factors that
interact to produce a suitable habitat for larval development (47). The distribution of the famin is world wide.
Most of the species in the family are thermophilous and adapted to living in standing water, although sper'-c do
occur in cold habitats and in running water (47). £. riparius is a non-biting midge. The tubiculous larvae
frequently inhabits cutrophic lakes, ponds, aua streams and reportedly live in mud-bottom lit tor.-'I habitats to
depths up to 1.0 meter (59). Qualitative observations indicate larvae inhabit gravel, limestone, marl, plants, ana
silt (53). Ingersoll and Nelson (4) report C. riparius to have a wide tolerance of sediment grain size. Sediment
ranging from >90% silt- and day-size particles to 100% .md-size particles did not reduce larval surv'val Or
growth in the laboratory. Larvae of C. riparius larvae reportedly occur in the fbld at a temperature range
between 0°C to 33*C, pH range between 5 to 9, and at dissolve, oxygen concentrations as low as 1 mg/L (53).
.£. riparius tubes arc of the type characteristic of bottom-feeding chironomid larvae (59). Larvae frequently
extend their anterior ends outside of their tubes feeding on the sedirr-nt surface (59). Credland (60) reported C.
riparius will eat a variety of materials of the appropriate size.
X3.2.1 The biology of C. riparius facilitates labc-atorv r- •— -:nce larvae are tolerant of a w .e spectrum of
conditions and adults mate even when confined (55,58,60). lu. vde of C. riparius can be divided into three
distinct stages: (1) a larval stage, consisting of the 4 instars; (2) a pupal stage, and (3) an adult stage. Midge egg
masses hatch in " ir 3 days after deposition in water at 19-22°C. Larval growth occurs in four instars of about 4-
7 days each. Unw^.- optimal conditions larvae will pupate and emerge as adults after 15 to 21 days at 20°C.
Adults emerge from pupal cases over a period lasting several days. Males are easily distinguished from females
because males have large, plumose antennae and a mucn thinner abdomen with visible genitalia. Mating
behavior has been described by Credland (60).
X33 rH^jp'Tg Test Orgat"SI"s • Tbe fo11owing is a description of culturing procedures adapted from
Ingenoll and Nelson (4) and others (51,54,^,60). These procedures should not be considered definitive. What
works in one Laboratory sometimes works poorly in another laboratory. £. riparius can be reared in aquaria in
either stalk or flowing water with a 16:8 hour light:darkness photoperiod at 20-22° C, at about 50 foot-candles
(538 lux). For static cultures the water should be gently aerated and about 25-30 percent of the water '• lume
should be replaced weekly. Cultures should be maintained in an isolated area or room free of contamination and
excessive disturbances. Ingersoli and Nelson (4) recommend rearing £. riparius in 30 x 30 x 30-cm polyethylene
containers covered with nylon screen. Each culture chamber contains 3 L of culture water. At least three egg
cases should be osod to start a new culture. To start a culture, 200-300 mg of ground cereal leaves 9 is added to
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
the culture chamber, additionally, green algae (Selenastn'm cflprjcprniJtum) (**1) ^ added ad libitum to maintain
a growth of algae in the water column and on the bottom of the culture chamber. Cultures should be fed about
3 mL of a suspension of commercial dog treats 12 (62) daily. This suspension should be prepared by heating and
melting 15 g of dog treats 12 in ISO mL of culture water. After refrigeration, the oily layer which forms on the
surface should be removed. The rest should be used to feed the cultures. This suspension contains about 100 mg
dry solid/mL. Overfeeding will lead to the growth of fungus in the aquaria and will necessitate more frequent
water changes. To obtain egg cases and larvae, adults should be left in the culture chamber to mate and deposit
eggs. Egg cases adhere to the side of the culture chamber and can be removed with a sharp blade. These egg
masses can then be placed in individual 100 mL beakers containing 50 mL of culture water; hatching should start
in about 3 days at 20° C. While removal of adults by aspiration into a 250 mL flask before mating works well
with £. tenfai^ (see Appendix X2), this procedure has not been successful with £. riparius.
X3.4 Brood Stock - Brood stock can be obtained from the wild, another laboratory or a commercial source.
When midges are brought into the laboratory, they should be acclimated to the culture water by gradually
changing the water in the culture chamber from the water in which they were transported to 100% culture water.
Midges should be acclimated to the test temperature by changing the water temperature at a rate not to exceed
2°C within 24 h, until the desired temperature is reached. Brood stock should be cultured so they are not
unnecessarily stressed. To maintain midges in good health and avoid unnecessary stress, crowding and rapid
changes in temperature and water quality characteristics should be avoided.
X3-5 Age - Tests with £. riparius can be started with either larvae less than 24-h old (4) or with three day
old larvae (56,57). Freshly laid midge egg cases can be transferred from the culture into individual 100 mL
beakers containing 50 mL of culture water. At 20°C larvae should begin to hatch within 3 days. Larvae must be
collected from at least three separate egg cases to start a sediment toxicity test.
X3.6 Handling - Midges should be handled as little as possible. When handling is necessary, it should be
done as gently, carefully, and quickly as possible, so that the midges are not unnecessarily stressed. First instar
midges should be transferred with a 2 mm inner diameter glass pipet (eye dropper). Older larvae should be
transferred with a 7 mm inner diameter glass pipet. Midges should be introduced into solutions beneath the air-
water interface. Any midges that touch dry surfaces, are dropped, or injured during handling should be
discarded.
X3.7 Afft'Tflflftfll - If the midges are cultured in water different from the overlying water or temperature, an
acclimation process is necessary. The water acclimation process used by Ingersoll and Nelson (4) is to first place
animals for 2h in a 50:50 mixture of culture water to overlying water, then for 2 h in a 25:75 mixture of culture
water to overlying water, followed by a transfer into 100% overlying water. At this stage the midges are
considered acclimated to the overlying water and should be ready for immediate use. Midges should be
randomly selected from the acclimation water with a pipette and placed into counting beakers (for example, 30-
mL) that can be floated in the test chambers before the midges are introduced into the exposure system.
X3.8 Toxicitv Test Specifications
12 Dog Kisses, The Hartz Mountain Corporation, Harrison, NJ 07029-9987.
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X3.8.1 Experimental PgSJgn • Decisions concerning the various aspects of experimental design, such as the
number of treatments, number of test chambers and midges per treatment, and water quality characteristics,
should be based on the purpose of the test and the type of procedure that is to be used to calculate results.
Ingersoll and Nelson (4) recommend using 50 JQ. riparius in each 1-L exposure beaker containing 200 mL of
sediment and 800 mL of overlying water in either static or flow-through testing. Lee (57) recommends using 13-
L glass aquaria containing 130 C. riparius larvae, 2 L of sediment and 11 L of overlying water in static tests.
Tests with C. riparius have been conducted at 20-22°C (4,5647)- Cooler test temperatures might reduce the
growth of fungus on the sediment surface. Duration of the test can range from a _<.10 day test to > 10 days and
continuing up to 30 days (4,56,57). Larval survival, growth, or adult emergence can be monitored as biological
endpoints.
X3.8.2 Static and Flow-through Tests - Ingersoll and Nelson (4) recommend that borosilicate glass 1-L
beakers can be used to expose the £. riparius to the test material, in either static or flow-through tests. For the
static tests, cover watch glasses may be used, such that an aeration line fits through the beaker pour spout and
the cover. For flow-through testing, Ingersoll and Nelson (4) suggest using a 4 x 13 cm notch cut in the lip of
the 1-L beaker. The notch should be covered with 033 mm U.S. Standard sieve size #50 screen, either made of
stainless steel or polyethylene, using a silicone adhesive to attach the screen to the beaker. For 24 hours after
hatching, first instar midge larvae are often planktonic (55). Pittinger et al. (56) suggest not running water
through the diluter for at least 24 hours after larvae are added to the test chambers. This will allow time for
larvae to settle onto the sediment surface.
X3.83 Initiation of a Test - Sediments are homogenized and placed hi the test chambers the day before
addition of test organisms (Day -1). Test chambers are then covered and overlying water is aerated overnight.
The test begins when midges are introduced to the test chambers (Day 0). Ingersoll and Nelson (4) start
sediment toxicity tests with 50 first instar £. riparius larvae per 1-L test chamber. Pittinger et aL (56) and Lee
(57) suggest starting tests with 3 day old larvae (130 larvae per 13-L chamber (57)). It is recommended that
flow-through and static tests might need to be started on different days to assure that sufficient time is available
to complete all tasks. Test chambers should be inspected <2 hours after midges are introduced to insure that
animals are not trapped in the surface tension of the water. These "floaters" do not survive well and should be
replaced with healthy animals.
X3.8.4 Feeding - Lee (57) recommends feeding animals in a static system 200 mg fish food flakes 10 every
other day to each 13-L test chamber containing 130 larvae. Pittinger et al. (56) suggest feeding animals in a
static renewal system with trout food 1S, dehydrated cereal leaves 9 (5:1 w/w) and commercial dog treats 12 daily
to each test chamber containing 20 larvae. In flow-through and static toxicity tests, Ingersoll and Nelson (4) feed
50 £. riparius larvae in each 1-L test chamber a combination of ground cereal leaves 9 (suspended in water), a
green algae ($. capricornutum) and commercial dog treats 12. In flow-through sediment toxicity tests, 75 mg of
ground cereal leaves 9, 30 mg of dog treats " and 6 x 107 S capricflmviflim algal cells should be added to each 1-
L test chamber the day test starts (day 0). From Day 1 to Day 6 of the test, 15 mg of ground cereal leaves 9
should be added to each test chamber; from Day 1 to Day 12, 30 mg of dog treats " should be added to each
test chamber and from Day 13 to the end of the test, 15 mg of dog treats 12 should be added to each test
13 Purina Trout Chow, Purina Mills Imx, 1401 S. Hanley, St Louis, MO 63144
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
chamber; 6 x 107 £. ragnfflrnufttm algal cells should be added to each test chamber daily. In static sediment
toxicity tests, 10 mg of ground cereal leaves 9, 10 mg of dog treats 12 and 3 x 107 5- capricornutum algal cells
should be added to each 1-L test chamber on Day 0. From Day 1 to Day 6 of the test, 10 mg of ground cereal
leaves 9 and 3 x 107 algal cells should be added to each 1-L test chamber; for the first two weeks of the test, 10
mg of dog treats 12 should be added to each test chamber each Monday, Wednesday, and Friday and for the rest
of the test 5 mg of dog treats 12 should be added to each test chamber each Monday, Wednesday and Friday,
from Day 7 until the end of the test 3 x 107 algal cells should be added to each test chamber each Monday,
Wednesday and Friday. Lower feeding levels for flow-through tests might be used for £. riparius daily: 6 x 107
algal cells, 10 mg dog treats 1Z, and 10 mg ground cereal leaves 9 on Day 0 - 6. If food collects
on the sediment, a fungal or bacterial growth might start on the surface of the sediment, in which case feeding
should be suspended for one or more days. A drop in dissolved oxygen to 40% saturation might indicate that all
of the food added in the water is not being consumed such that feeding should be suspended for the amount of
time necessary to increase the dissolved oxygen concentration (4).
X3.8.5 Biological Data - Several endpoints can be monitored in midge sediment toxicity tests. During the
test, emergence of larvae from the test sediment can be monitored. Additionally, data on larval survival, growth,
and adult emergence can be obtained.
X3.8.5.1 Larval survival and growth can be assessed by ending the tests on Day 10 to Day 14 when larvae
have reached the 3rd or 4th instar (4,25,45). At this time, larvae should be removed from sediment using a #35
(500 Jim) U.S. Standard size sieve (4). The midges should be rinsed from the sieve into collecting pans and
pipeted from the rinse water. Growth determination using dry weight (dried at 60°C to a constant weight) is
preferable to length. Growth can also be estimated by measuring head capsule width, and also used to determine
instar development
X3.8.5.2 Ingersoll and Nelson (4), Pittinger et al. (56) and Lee (57) recommend conducting C. riparius
sediment toxicity tests until the larvae pupate and emerge as adults. Cast pupal skins left by emerging adult £.
riparius should be removed and recorded daily. These pupal skins remain on the water surface for over 24 hours
after the emergence of the adult. The test should be ended after the animals have been exposed for up to 30
days, when about 70-95% of the control larvae should have completed metamorphosis into the adult form.
Endpoints calculated in these adult emergence tests can include: (1) percent emergence, (2) mean emergence
time, or (3) day to first emergence. Egg hatching studies may also be conducted by covering the test chambers
and confining the adults. Adults will emerge and lay eggs in these chambers. These egg masses can then be
used to estimate effects of exposure on either the number of eggs produced or hatched.
X3.8.53 A C. riparius sediment toxicity test, independent of duration, is unacceptable if the average survival
in any negative control chamber is less than 70% (see Section 15, Acceptability of Test). (Note: a low percent
adult emergence might not be the result of low survival; larvae or pupae might not have completed development).
33
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53. Curry, LL. 1962. A survey of environmental requirements for the midge. (Diptera: Tendipedidae). In
Biological Problems in Water Pollution, pp. 127-14L US. Pub. Health Serv. Publ. 999-WP-25, Cincinnati,
OH, 376 p.
54. Batac-Catalan, Z. and D.S. White. Creating and Maintaining Cultures of Chironomus tg'Uanfi (Diptera:
Chironomidae). Entomological News. Vol. 93. pp. 54-58. 1982. Yount, J. 1966. A method for rearing large
numbers of pond midge larvae, with estimates of productivity and standing crop. American. Midland
Naturalist 76:230-238; McLarney, W.O., S. Henderson and MJM. Sherman. 1974. A new method for culturing
Fabricius larvae using burlap substrate in fertilized pools. Aquaculture 4:267-276;
Nebeker, A.V., M.A. Cairns, and CM. Wise. Relative Sensitivity nf rhirnqomus tentans Life Stages to
Copper. Environmental Contamination and Toxicology. Vol. 3. pp. 151-158. 1984.
55. Davies, BJL 1976. The Dispersal of Chironomidae Larvae: A Review. J. ent. Soc sth. Afr. 39:39-59.
56. Pittinger, CA^ DM. Weltering and JA. Masters. Bioavailabiliry of sediment-sorbed and soluble sufactants to
(Midge). Environ. ToxicoL Chem. 8(11), 1989.
57. Lee, CM. 1986. Toxicity of dihard-tallow dimethyl ammonia chloride. Tenside Detergents 23:196-199.
58. Powlesland, C. and J. George. 1986. Acute and chronic toxicity of nickel to larvae of Chironomus riparius
(Meigen). Environ. PolL 42:47-64; Wegner, G.S. and R.W. Hamilton. 1976. Effect of calcium sulfide on
ppaTIUS (Diptera: Chironomidae) egg hatchability. Environ. EntomoL 5:256-258; Williams, KA.,
Oecologia 70-362-366.
D.WJ. Green, D. Pascoe, and D.E. Gower. 1986. The acute toxicity of cad mum to different larval stages of
(Diptera: Chironomidae) and its ecological significance for pollution regulation.
36
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop ASTM E 1383 (in press)
59. Rasmussen, JJJ. 1984. The Life-history, Distribution, and Production of Chironomus riparius and
Glvptotendipes paripes in a Prairie Pond. Hydrobiologia 119:65-72.
60. Credland, PJ7. 1973. A new method for establishing a permanent laboratory culture of Chironomus riparius
Meigen (Diptere Chironomidae). Freshwater Biology 3:45-51;
61. Miller, WJL, J.C. Greene, and T. Shiroyama. 1978. The -^fclenastry.^ capricorniitym assay bottle test.
Experimental Design, Application, and Data Interpretadon ProtocoL EPA-600/9-78-018; Interim Procedures
for conducting the, n^phjiigi magna toricity assay. Environmental Research Laboratory, Duluth, MN 55804
and Environmental Monitoring Systems Laboratory, Las Vegas NV 89114. Office of Research and
Development U.S. EPA. February 1984.
62. Biever, K.D. A rearing technique for the colonization of chironomid midges. Ann^lg of the Entomological
Society of America. Vol. 58. DD. 135-136. 1965.
37
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Nelson, Coyle and Burton MPCB 1991: Sediment Workshop Fathead Sediment Testing
Contact: G. Allen Burton
Biological Sciences Dept.
Wright State University
Dayton, OH 45435
FATHEAD MINNOW WHOLE SEDIMENT TOXICITY TESTING
JP. promelas culturing and test conditions are similar to the effluent test method guidance of the
USEPA. Larvae (40 per treatment, 10 per beaker) less than 24h old are used in toxitity tests. The larvae
are randomly added to 600 mL test beakers containing 625 mL sediment and 250 mL overlying water. In
tests exceeding 48h exposure periods, the larvae are fed brine shrimp nauplii (0.1 mL, ~ 1050-1500
organisms) twice daily. Overlying waters are siphoned (80%) daily and replaced with fresh reconstituted,
after removing larvae, the fish are weighed at time zero (subsample) and after 7 days in chronic tests using
growth as the endpoint. Fish are removed on Day 7, placed in pre-weighed aluminum pans and dried at
105°C for 2 to 24 h. Dried larvae are weighed in groups of 10 on a Mettler balance. For the embryo-larval
assays, test chambers (600 mL beakers) receive 50 mL of sediment and 200 mL of site water to provide a 1:4
ratio of sediment to water by volume. The water is slowly added to prevent sediment resuspension. Ten
freshly-spawned embryos (less than 24h old) are added to each test chamber using a large bore pipette. Four
replicate chambers are run per test concentration. A set of control chambers contain no sediment is run in
quadruplicate concurrently with the exposure beakers. The water in the test beakers is gently and
continuously aerated for the duration of the test. Approximately 80% of the dilution water in each test
chamber is siphoned and renewed daily during the test period. Dead organisms are counted and removed
daily. Organisms are considered dead when they become opaque and white. Test organisms are not fed
during the test period. The temperature of the dilution water is maintained at 25"C, and a 16:8h light/dark
photoperiod is used. The temperature, pH, and dissolved oxygen content is monitored daily, while the
hardness, alkalinity, and conductivity of the test solution is measured at the beginning and end of the test, at
a minimum. At the end of the test period, the surviving test organisms are counted, removed, and placed
into a preservative until the larvae can be examined microscopically for terata and the lengths measured.
The endpoints of test measured are 7 d survival, growth (as measured by length), and percent hatch.
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Contact G. Allen Burton, Jr.
Wright State University
Biological Sciences Dept
Dayton, OH 45435
Tel. (513) 873-2201
FAX (513) 873-4106
Draft No. 3
ASTME1383. ANNEX X4. Daphnia and Ceriodaphnia sp.
X4.1 Significance - Daphnia magna and Ceriodaphnia dubia have many
desirable characteristics as toxicity test organisms. They are easily cultured in the
laboratory, have a short generation time, survival and reproduction data can be
obtained in toxicity tests, and a large data base has developed regarding their
sensitivity to toxicants. Nebeker et aL (1), Prater and Anderson (2), Giesy et al. (3),
Malueg et al. (4) and Burton et al. (5) and others (6-15) have successfully used
cladocerans in sediment testing and have shown them to be sensitive indicators of the
presence of contaminants associated with sediments.
In whole sediment toxicity tests, dadocera behave as nonselective epibenthic
zooplankton. The organisms are frequently observed on the sediment surface and are
likely exposed to both water soluble and particulate bound contaminants in overlying
water and surface sediments. These routes of exposure do not, however, mimic those
of infaunal benthic invertebrates, which are exposed directly to sediment and
interstitial water. One of the most important reasons for using cladocerans as toxicity
test organisms is their importance in the food web of some systems (16-18). Also
these assays have been useful at discriminating sediment contamination and allowing
comparisons of relative sediment toxicity. Because they are not benthic organisms,
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their responses may not be indicative of in situ benthic community effects.
X4.2 Life History and Life Cycle - Pennak (18) recognizes four distinct periods
in the life history of a dadoceraru 1) eg£ 2) juvenile, 3) adolescent, and 4) adult
Unstressed populations consist almost exclusively of females producing diploid
parthenogenetic eggs which develop into female young. An adult Ceriodaphnia can
produce from 4 to 15 parthenogenetic eggs in each brood whereas Daphnia can
produce 5 to 25 or more eggs (19). Pennak (18) indicates that when a dutch of eggs
is released into the brood chamber, segmentation begins promptly; the first juvenile
instar is released into the surrounding water in approximately two days. There are
only a few juvenile instars and the greatest growth occurs during these stages. The
adolescent period is a single instar between the last juvenile instar and the first adult
instar during which the first dutch of eggs reaches full development in the ovary. At
the dose of the adolescent instar, the animal molts and the first dutch of eggs is
released into the brook chamber, while a second dutdi is developing in the ovary.
At the dose of each adult instar, four successive events occur 1) the young are
released from the brood chamber to the outside environment, 2) molting occurs, with
3) an increase in size, and 4) mere is release of a new dutch of eggs into the brood
chamber.
When populations are under stress (e.g., low oxygen, crowding, starvation),
males are produced from diploid parthenogenetic eggs. When males appear, females
produce haploid eggs which require fertilization. Following fertilization, the eggs are
endosed by the ephippium and shed at the next molt The embryos lie dormant until
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suitable conditions arise upon which they become females producing diploid
parthenogenetic eggs (20).
X4.3 Obtaining Test Organisms - The following culture procedures are
adapted from Knight and Waller (21), while other appropriate methods include the
U.S. Environmental Protection Agency (22,23) ASTM E729 and E1295. Following
Knight and Waller's (21) methodology, D. magna and Ceriodaphnia dubia can be
cultured in reconstituted hard water (160-180 mg/L CaCOa) and fed a daily diet of a
vitamin enriched Selenastrum capricornutum suspension. Cultures are maintained at
25°C +. 1°C with a lighbdark cycle of 16:8 hours provided by overhead fluorescent
lighting covered with opaque plastic to reduce light intensity to less than 20 lux. D.
magna mass cultures are started by placing 10 neonates (less than 24 hours old) into
one liter beakers containing 500 ml reconstituted hard water and 12 ml
(approximately 240,000 algal cells/ml culture water) of S. capricornutum feeding
suspension. Cultures are fed 12 ml initially and on day one, 25 mis (500,000 cells/ml
culture water) on day two through four, and 25 to 50 mis (100,000 cells/ml culture
water) on day five and thereafter. Using this culture method, D. magna typically will
have first broods between days 6 and 8 with successive broods hatching every 36-48
hours thereafter. On days when hatches occur and young are not needed, adults are
transferred to dean one liter beakers containing 300 ml hard water, 200 ml old
culture water, and 50 ml of food. When young are needed for testing, the adults are
isolated the night before by placing each adult into a separate 100 ml beaker
containing 100 ml reconstituted hard water and 3 ml feeding suspension. Isolating
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adults into smaller beakers allows one to easily remove individual young for testing.
Neither first brood young nor young from females older than two weeks are used in
toxicity testing or initiating new cultures. The Selenastrum capricomutum feeding
suspension may also be supplemented with an approximate 6% by volume addition
of a Cerophyl® preparation to the algal feeding suspension (Waller, personal
communication). C dubia mass cultures can be initiated by placing 20 neonates (less
than 12 h old) into a 600 ml beaker containing 360 ml reconstituted hard water and
12 ml of S. capricomutum feeding suspension. Cultures are fed 12 ml initially and
on days one and two, and then 18 mis thereafter. When three distinct sizes are noted
(generally day 6) then the largest organisms are isolated in 100 ml beakers containing
60 ml of hard water and 2 ml feeding suspension. Less than 12 h old neonates from
the next brood (third brood) are used in toxicity testing and initiating new mass
cultures. Generally, first broods are produced on day four, second brood on day 5
and third brood on day 7. Isolated females generally produce between 10 and 16
neonates on their third brood (21).
The U.S. Environmental Protection Agency (23) recommends culturing D.
magna in reconstituted hard water at 20°C with ambient light intensity of 50-100 ft c
(10-20 uE/mVs, or 538-1076 lux), and a light:dark cycle of 16:8 hours. Culture
vessels can be 3 L glass beakers containing 2.75 L reconstituted hard water and 30 D.
magna. The D. magna can be fed on a daily diet of S. capricomutum (100,000 algal
cells/ml culture water) or fed three timers a week on a feeding suspension consisting
of trout chow, alfalfa and yeast (TCY) (1.5 ml TCY/1000 ml culture water). This
should supply approximately 300 young per week.
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The U.S. Environmental Protection Agency (22) procedures for Ceriodaphnia
cultures are as follows. Ceriodaphnia are cultured in moderately hard water (80-90
mg/L CaCOj) at 25°C ± 1°C and receive a lighfcdark cycle of 16:8 hours. Mass
cultures are maintained as "backup" organism reservoirs and individual organisms
are cultured as the source of neonates for toxicity tests. Mass cultures can be
initiated in 2-3 L beakers filled to three-fourths capacity with moderately hard water
and 40-50 neonates per liter of medium. The stocked organisms should be
transferred to fresh culture media twice weekly for two weeks. At each renewal, the
adults are counted and the offspring and old medium discarded. The adults are
discarded after two weeks and new mass cultures initiated with neonates. Mass
cultures are fed daily at the rate of 7 ml of a yeast, Cerophyl, trout chow food
preparation (YCT)' and 7 ml of J>. capricornutum concentrate (3.0-3.5 x 107 cells/ml).
Individual C dubia cultures are maintained in 30 ml plastic cups or beakers
containing 15 ml of culture media. Cultures are fed daily at the rate of 0.1 m YCT
and 0.1 ml algal concentrate per 15 ml media and are transferred to fresh media at
least three times a week. Adults are used as sources of neonates until 14 days of age.
Cultures properly maintained should produce at least 15 young per adult in three
broods (seven days or less). Goulden and Henry (19) list two other fresh water algal
species which can be used for cladoceran food: 1) Ankistrodesmus falcatus, and 2)
Chlamydomonas reinhardtii. Winner (24) discusses the effects of four diets
fChlamvdomonas reinhardtii. Selenastrum capricornutum. yeast-trout chow-Cerophyl
(YTQ, and YTC plus S. capricomutuml and two reconstituted waters on the vitality
of five to six lifespan generations of C dubia. His results indicate that healthy
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populations can be maintained in reconstituted hard water containing only four salts
as long as the food is nutritionally adequate and the water is reconstituted from an
ultrapure base water.
X4.4 Brood Stock - D. magna and C dubia starter cultures can be obtained
from the Aquatic Biology Branch, Environmental Monitoring Systems Laboratory,
USEPA, 3411 Church Street, Newtown, OH 45244. Animals received from an outside
source should be acclimated gradually to new culture media over a period of 1-2
days.
X4.5 Background - The various decisions concerning experimental design,
such as number of test chambers, number of treatments, animals per treatment and
water quality characteristics, should be based on the purpose of me test and the
procedure used to calculate results. See ASTM E729, E1295, E1297, and the preceding
guide text for guidance. Nebeker et aL (25) recommended conducting 48 h sediment
static tests in duplicate using 1 L beakers containing 200 ml of sediment and 800 ml
of water (1:4 ratio). The sediment is allowed to settle overnight, followed by gentle
aeration of overlying water for 30 minutes before introducing 15 D. magna per
replicate. Malueg et al (4) conducted recirculating sediment toxicity tests in a
modified recycling device described by Prater and Anderson (2). The test chamber
(23 cm long x 6.4 on wide x 16 cm high) was positioned on a Plexiglass plate over
two 4 - L jars. Twenty D. magna were placed in a vessel in the water column and 5
Hexagenia added to chamber sediment Three to six replicates were used for each
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control and test sediment Seven day (three brood) toxicity tests for aqueous media
using dadocerans have been conducted (1,26,27) and variations of these methods
used to assess sediment toxicity (1,28).
X4.6 Handling - The dadocerans are delicate and should be handled as
carefully and little as possible. They are transferred with a 5 mm bore pipet and
released slowly beneath the water surface.
X4.7 Experimental Design for Acute Toxicitv Tests - Sediments may be mixed,
if appropriate for the study, by mixing with either a large plastic paddle, magnetic
stirring bar or shaker table, before allocating to test chambers. See ASTM 1297 and
1391 for guidance. Whole sediment assays use a 1:4 ratio of sediment to water.
Acute toxicity tests are conducted in triplicate using 250 or 100 ml beakers to which
30 ml of sediment (by weight) and 120 ml of reconstituted or site water are added
(for 250 ml beakers). The weight of 30 ml of sediment is determined by initially
calculating the average wet weight (grams) of five, 5 ml aliquots of sediment
obtained using a 10 cc syringe. The average weight of 5 ml is divided by five to
obtain the weight of 1 ml of sediment The weight of 1 ml is multiplied by 30 ml to
obtain the number of grams to be weighed into each test beaker. When a syringe
cannot be used to dispense sediments, sediment weight is used rather than volume,
weighing 30 grams (wet weight) into each test beaker. In addition, sediment dry
weights are determined by weighing triplicate 3-5 ml aliquots of wet sediment;
drying at 100-105°C for 24 hours and then reweighing the sediment Percent dry
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weight is calculated by dividing the dry sediment weight (grams) by the wet weight
and multiplying by 100. Grams of dry weight per ml of wet sediment is determined
by dividing the dry weight by the ml of wet sediment Overlying water is gently
added to each beaker, minimizing sediment resuspension. After a 1 to 2 hr settling
period, ten test organisms are randomly added to each beaker. Test chambers should
be inspected less than 2 h after the addition of test organisms to check for any
"floaters." "Floaters" may not survive and are subjected to a different exposure, thus
can be removed and replaced within the first two hours. Floating may be caused by
the sediment sample and may be considered a treatment effect in some cases.
However, responses tend to be variable and are seldom dose proportional. Surface
films which entrap D. magna can be reduced by wiping the surface with cellulose
filter paper prior to organism addition.
X4.8 Experimental Design for Short-term Chronic Toxicirv Tests. Test
initiation, test conditions and monitoring are as described in Section X4.7 and X4.9
with the following exceptions, and basically follow standard methods (22, ASTM
E1295). Tests are conducted in 30 ml beakers using 5 ml (or grams) sediment and 20
ml overlying water in replicates of ten. One organism (D. magna less than 24 hr old
or C dubia less than 6 hr old) is randomly added to each beaker, after the settling
period. At each 24 hr test interval, the adult is removed and placed in a beaker
containing the control water, young are counted and discarded, and physicochemical
measures made. Approximately 15 ml of overlying water is suctioned off and gently
renewed. The culturing food (such as YCT or algal-Cerophyl® mixture) is then
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added (0.1 ml) to each beaker. After feeding, the adult organism is returned to the
test beaker. The test is terminated at 7 days and /or when at least 60% of the controls
have produced their third brood.
X4.9 Monitoring Data - Test conditions and monitoring should follow
standard methods (22,23). Test beakers are maintained at 25 ± 1°C and receive a
16:8 h lighfcdark cycle (20 lux). Dissolved oxygen and temperature are monitored at
0, 24 and 48 h. Dissolved oxygen should not be allowed to drop below 40%
saturation. If it does, gentle bubbling should be used until adequate saturation is
attained. The pH, hardness and alkalinity are monitored at 0 and 48 h. Survival
numbers were recorded at 24 and 48 h. Death of a test animal is judged as a result
of observing no movement upon gentle prodding. Tests are considered valid when
control mortality is £ 10% (23). Control treatments consist of reconstituted water or
reference site water, and a control and/or reference sediment with the overlying test
water (reconstituted or reference site). See the preceding guide text for additional
guidance on sediment characterization, controls, references, and data analyses.
The 7-day survival and reproduction test requires the daily counting of adult
survivors and young production. Dissolved oxygen, temperature, and pH should be
measured daily, before renewing overlying waters on two to three beakers in each
treatment and control Alkalinity and hardness are measured at test initiation and
termination. For the test results to be acceptable controls must have 80% survival
with C. dubia controls averaging 15 young and D. maena averaging 60 young per
surviving female (22^6).
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Literature Cited
1. Nebeker, A.V., MA. Cairns, J.H. Gakstatter, K.W. Malueg, G.S. Schuytema, and
D.F. Krawczyk. 1984. Biological methods for determining toxirity of
contaminated freshwater sediments to invertebrates. Environ. Toxicol.
Chem. 3: 617-630.
2. Prater, B.L and MA. Anderson. 1977. A % hour sediment bioassay of
Duluth and Superior Harbor Basins (Minnesota) using Hexagenia
limbata. Asellus communis. Daphnia maena, and Pimephales promelas
as test organisms. Bull. Environ. Contain. Toxicol. 18:159-169.
3. Giesy, J.P., CR. Rosiu, and R.L. Graney. 1990. Benthic invertebrate bioassays
with toxic sediment and pore water. Environ. Toxicol. Chem. 9:233-248.
4. Malueg, K.W., G.S. Schuytema, J.H. Gakstatter, and DJ. Krawczyk. 1983.
Effect of Hexagenia on Daphnia response in sediment toxicity tests.
Environ. Toxicol. Chem. 2: 73-82.
5. Burton, G.A., Jr., B.L Stemmer, K.L. Winks, P.E. Ross, and LC Burnett. 1989.
A multitrophic level evaluation of sediment toxicity in Waukegan and
Indiana Harbors. Environ. ToxicoL Chem. 8:1057-1066.
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toxicity changes in the presence of sediment Bull. Environ. Contain.
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13. Giesy, J.P., R.L. Graney, J.L. Newsted, CJ. Rosiu, A. Benda, R.G. Kreis, Jr. and
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16. Mount, DJ. and TJ. Norberg. 1984. A seven day life-cycle cladoceran toxicity
test Environ. Toxicol. Chem. 3: 425-434.
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18. Pennak, R.W. 1978. Freshwater invertebrates of the United States. 2nd ed.
John Wiley and Sons, New York, NY.
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20. Lawrence, S.G. (ed.). 1981. Manual for the culture of selected freshwater
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22. U.S. Environmental Protection Agency. 1985. Short-term methods for
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23. U.S. Environmental Protection. Agency. 1985. Methods for measuring the
acute toxicity of effluents to freshwater and marine organisms. EPA
600/4-85/013. Cincinnati, OR
24. Winner, R.W. 1989. Multigeneration life-span tests of the nutritional adequacy
of several diets and culture waters for Ceriodaphnia dubia. Environ.
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25. Nebeker, Alan V., S.T. Onjukka, MA. Cairns, and DJ. Krawczyk. 1986.
Survival of Daphnia magna and Hyalella azteca in cadmium spiked
water and sediment Environ. Toxicol. Chem. 5: 933-938.
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magna and Ceriodaphnia dubia toxicity tests for cadmium and sodium
pentachlorophenate. Environ. Toxicol. Chem. 7:153-156.
27. Mount, D.I. and T.J. Norberg-King. 1984. A seven-day life cycle dadoceran
toxicity test Environ. Toxicol. Chem. 3: 425-434.
28. Burton, G.A., Jr., L. Burnett, M. Henry, S. Klaine, P. Landrum, and M. Swift
1990. A multi-assay comparison of sediment toxicity at three "Areas of
Concern," Abstr. Annu. Meet Soc. Environ. Toxicol. Chem., Arlington,
VA. No, 213, p. 53.
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